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Full text of "soil analysis"

R. Margesin F.Schinner (Eds.) 



Soil Biology 



Series Editor: Ajit Varma 



5 



Volumes published in the series 



Volume 1 

A. Singh, O.P. Ward (Eds.) 

Applied Bioremediation and Phytoremediation 

2004 

Volume 2 

A. Singh, O.P. Ward (Eds.) 

BlODEGRADATION AND BlOREMEDIATION 

2004 

Volume 3 

E Buscot, A. Varma (Eds.) 

Microorganisms in Soils: Roles in Genesis and Functions 

2005 

Volume 4 

S. Declerck, D.-G. Strullu, J.A. Fortin (Eds.) 

In Vitro Culture of Mycorrhizas 

2005 



Rosa Margesin 
Franz Schinner (Eds.) 



Manual for Soil Analysis 
Monitoring and Assessing 
Soil Bioremediation 



With 3 1 Figures 



4y Spri 



ringer 



Prof. Dr. Rosa Margesin 

Leopold Franzens University 

Institute of Microbiology 

Technikerstr. 25 

A-6020 Innsbruck 

Austria 

e-mail: Rosa.Margesin@uibk.ac.at 

Prof. Dr. Franz Schinner 

Leopold Franzens University 

Institute of Microbiology 

Technikerstr. 25 

A-6020 Innsbruck 

Austria 

e-mail: Franz.Schinner@uibk.ac.at 



Library of Congress Control Number: 2005926091 

ISSN 1613-3382 

ISBN-10 3-540-25346-7 Springer Berlin Heidelberg New York 

ISBN-13 978-3-540-25346-4 Springer Berlin Heidelberg New York 



This work is subject to copyright. All rights reserved, whether the whole or part of the 
material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, 
recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data 
banks. Duplication of this publication or parts thereof is permitted only under the provisions 
of the German Copyright Law of September 9, 1965, in its current version, and permission 
for use must always be obtained from Springer. Violations are liable for prosecution under 
the German Copyright Law. 

Springer is a part of Springer Science + Business Media 

springeronline.com 

© Springer- Verlag Berlin Heidelberg 2005 
Printed in Germany 

The use of general descriptive names, registered names, trademarks, etc. in this publication 
does not imply, even in the absence of a specific statement, that such names are exempt 
from the relevant protective laws and regulations and therefore free for general use. 

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Typesetting and production: LE-TgX Jelonek, Schmidt & Vockler GbR, Leipzig, Germany 

31/3150-YL - 5 4 3 2 1 - Printed on acid-free paper 



Preface 



The increasing use of soil bioremediation technologies requires new con- 
cepts and methods to assess the feasibility of a remediation technology 
and to monitor the success of the treatment. The knowledge of the reac- 
tion of the soil microflora to contamination facilitates the optimization of 
biodegradation. Manual of Soil Analysis - Monitoring and Assessing Soil 
Bioremediation differs from other books on soil analysis in that the moni- 
toring and assessing of soil bioremediation are the central themes. 

In this comprehensive laboratory manual, sampling, pretreatment and 
storage of soil, feasibility studies for soil bioremediation, and the most 
important methods to analyze physical, chemical, and biological soil pa- 
rameters are presented. Chapters written by experts for those involved in 
research, teaching, and routine analyses outline molecular and immunolog- 
ical techniques, the use of conserved internal markers, radiorespirometry, 
bioreporter technology, the interpretation of fatty acid profiles, soil mi- 
crobial and enzymatic methods, and the assessment of ecotoxicity using 
bioassays. Particular emphasis has been placed on the comprehensible and 
complete description of the experimental procedures. The broad spectrum 
of modern soil biological methods provides an excellent complementation 
of traditional soil investigation and characterization. Our book, however, 
does not claim to present all modern methods available, it rather contains 
a selection of the most suitable methods for investigating contaminated 
soil. More biological methods can be found in our volume Methods in Soil 
Biology (Schinner, Ohlinger, Kandeler and Margesin 1996, Springer). 

We are most grateful to the authors for their excellent contributions and 
to Springer, especially to Dr. Jutta Lindenborn and Dr. Dieter Czeschlik, for 
continuous support and cooperation. We also thank Dr. Ajit Varma for the 
possibility to publish this book in the Soil Biology Series. 

Innsbruck, Austria, Rosa Margesin 

January 2005 and Franz Schinner 



Contents 



1 Soil Sampling and Storage 1 

Andreas Paetz, Berndt-Michael Wilke 

1.1 Objective of Soil Sampling 1 

1.1.1 Principal Objectives 1 

1.1.2 Specific Objectives 4 

1.2 Selection of Sampling Technique 6 

1.3 Sampling Strategy 7 

1.3.1 General 7 

1.3.2 Preliminary Investigation 7 

1.3.3 Exploratory Investigation 9 

1.3.4 Main Site Investigation 9 

1.3.5 Samples and Sampling Points 10 

1.4 Sampling Methods 25 

1.4.1 General 25 

1.4.2 Type of Sample 25 

1.4.3 Undisturbed Samples 27 

1.4.4 Cross-Contamination 34 

1.4.5 Sampling Containers 34 

1.5 Pretreatment 37 

1.5.1 Chemical Analysis 37 

1.5.2 Physical Analysis 40 

1.5.3 Biological Analysis 40 

1.6 Storage of Samples 41 

1.6.1 General 41 

1 .6.2 Specific Considerations for Biological Parameters 42 

1.6.3 Preparing the Samples After Storage 44 

References 44 

2 Determination of Chemical and Physical Soil Properties 47 

Berndt-Michael Wilke 

2.1 Soil Dry Mass and Water Content 47 

2.2 Water-Holding Capacity 50 



VIII Contents 

2.3 Bulk Density - Total Porosity 52 

2.3.1 Core Method 52 

2.3.2 Excavation Method 54 

2.3.3 Clod Method 57 

2.4 Water Retention Characteristics - Pore Size Distribution 59 

2.4. 1 Determination of Soil Water Characteristics 

Using Sand, Kaolin, and Ceramic Suction Tables 62 

2.4.2 Determination of Soil Water Characteristics 

by Pressure Plate Extractor 65 

2.5 SoilpH 68 

2.6 Soil Organic Matter - Soil Organic Carbon 71 

2.6.1 Dry Combustion Method 72 

2.6.2 Loss On Ignition Method (LOI) 74 

2.7 Soil Nutrients: Total Nitrogen 76 

2.7. 1 Dry Combustion Method ("Elemental Analysis") 77 

2.7.2 Modified Kjeldahl Method 79 

2.8 Soil Nutrients: Inorganic Nitrogen 82 

2.8.1 Extraction 83 

2.8.2 Quantification of Nitrate Nitrogen 84 

2.8.3 Quantification of Ammonium Nitrogen 86 

2.9 Soil Nutrients: Phosphorus 87 

2.9.1 Extraction of Total Phosphorus 88 

2.9.2 Extraction of Labile Phosphorus 90 

2.9.3 Quantification of Phosphorus 91 

References 93 

3 Quantification of Soil Contamination 97 

Kirsten S. j0rgensen, Olli Jarvinen, Pirjo Sainio, Jani Salminen, 
Anna-Mari Suortti 

3.1 General Introduction 97 

3.2 Volatile Hydrocarbons 99 

3.3 Hydrocarbons in the Range C 10 to C 40 103 

3.4 Polyaromatic Hydrocarbons (PAHs) 109 

3.5 Heavy Metals 115 

References 118 

4 Immunotechniques as a Tool for Detection of Hydrocarbons 121 

Grazyna A. Plaza, Krzysztof Ulfig, Albert J. Tien 

4.1 RaPID Assay Test System 121 

4.2 EnviroGard Test System 126 

References 130 



Contents IX 

5 Feasibility Studies for Microbial Remediation 

of Hydrocarbon-Contaminated Soil 131 

Ajay Singh, Owen P. Ward, Ramesh C. Kuhad 

5.1 Introduction 131 

5.2 Determination of Biodegradation Potential 132 

5.2.1 Sampling and Soil Preparation 132 

5.2.2 Selective Microbial Enrichment 134 

5.2.3 Controls 135 

5.2.4 Soil Microcosms 136 

5.2.5 Slurry Bioreactors 137 

5.2.6 Land Treatment 139 

5.2.7 Composting 140 

5.2.8 Scale-Up 141 

5.3 Process Monitoring and Evaluation 142 

5.4 Bioaugmentation 143 

5.5 Effect of Surfactants 144 

5.5. 1 Screening of Microbial Cultures 

for Biosurfactant Production 145 

5.5.2 Effect of Biosurfactants 146 

5.5.3 Effect of Chemical Surfactants 146 

5.6 Optimization of Environmental Conditions 147 

5.7 Optimization of Nutritional Factors 148 

5.8 Conclusions 150 

References 151 

6 Feasibility Studies for Microbial Remediation 

of Metal-Contaminated Soil 155 

Franz Schinner, Thomas Klauser 

References 159 

7 Feasibility Studies for Phytoremediation 

of Metal-Contaminated Soil 161 

Aleksandra Sas-Nowosielska, Rafal Kucharski, Eugeniusz Malkowski 

7.1 Introduction 161 

7.2 Phytoextraction 161 

7.2.1 Treatability Study 162 

7.2.2 Full-Scale Application 166 

7.2.3 Conclusions 170 

7.3 Phytostabilization Potential for Soils Highly Contaminated 
with Lead, Cadmium and Zinc 171 

7.3.1 Evaluation of Site Contaminants 171 

7.3.2 Logistic Considerations 172 



X Contents 

7.3.3 Additives 172 

7.3.4 Plants 173 

7.3.5 Full-Scale Application 173 

7.3.6 Effectiveness of Technology 174 

7.3.7 Monitoring 174 

7.3.8 Conclusions 175 

References 176 

8 Quantification of Hydrocarbon Biodegradation 

Using Internal Markers 179 

Roger C. Prince, Gregory S. Douglas 

References 187 

9 Assessment of Hydrocarbon Biodegradation Potential 

Using Radiorespirometry 189 

Jon E. Lindstrom, Joan F. Braddock 

References 198 

10 Molecular Techniques for Monitoring 

and Assessing Soil Bioremediation 201 

Lyle G. Whyte, Charles W. Greer 

10.1 General Introduction 201 

10.2 Extraction and Purification 

of Nucleic Acids (DNA) from Soil 202 

10.3 Amplification of Catabolic Genotypes 

and 16SrDNA Genotypes by PCR 208 

10.4 DGGE Analysis Soil Microbial Communities 218 

10.5 Genomics in Environmental Microbiology 226 

References 228 

1 1 Bioreporter Technology for Monitoring Soil Bioremediation 233 

Steven Ripp 

11.1 General Introduction 233 

11.2 An Overview of Reporter Systems 

for Soil Bioremediation Application 235 

11.3 Single Point Measurements of Soil Contaminants 241 

1 1.4 Continuous On-Line Vapor Phase Sensing 

of Soil Contaminants 244 

1 1.5 Quantification of Soil-Borne lux-Tagged Microbial Popula- 
tions Using Most-Probable-Number (MPN) Analysis 247 

References 249 



Contents XI 

12 Interpretation of Fatty Acid Profiles of Soil Microorganisms 251 

David B. Hedrick, Aaron Peacock, David C. White 

12.1 Obtaining Fatty Acid Profiles from Soil Samples 251 

12.2 Transforming Fatty Acid Peak Areas 

to Total Microbial Biomass 252 

12.3 Calculation and Interpretation of Community Structure 254 

12.3.1 Standard Community Structure Method 254 

12.3.2 Custom Community Structure Methods 255 

12.3.3 Factor Analysis 255 

12.4 Calculation and Interpretation 

of Metabolic Stress Biomarkers 256 

12.5 Naming of Fatty Acids 257 

References 258 

13 Enumeration of Soil Microorganisms 261 

Julia Foght, Jackie Aislabie 

13.1 Sample Preparation and Dilution 261 

13.2 Direct (Microscopic) Enumeration 264 

13.3 Enumeration by Culture in Liquid Medium 

(Most Probable Number Technique) 268 

13.4 Enumeration by Culture on Solid Medium 

(Plate Count Technique) 272 

References 279 

14 Quantification of Soil Microbial Biomass 

by Fumigation-Extraction 281 

Rainer Georg Joergensen, Philip C. Brookes 

14.1 General Introduction 281 

14.2 Fumigation and Extraction 282 

14.3 Biomass C 284 

14.3.1 Biomass C by Dichromate Oxidation 284 

14.3.2 Biomass C by UV-Persulfate Oxidation 286 

14.3.3 Biomass C by Oven Oxidation 288 

14.4 BiomassN 289 

14.4.1 Ninhydrin-Reactive Nitrogen 289 

14.4.2 Total Nitrogen 292 

References 294 

15 Determination of Adenylates and Adenylate Energy Charge 297 

Rainer Georg Joergensen, Markus Raubuch 

References 302 



XII Contents 

16 Determination of Aerobic N-Mineralization 303 

Rainer Georg Joergensen 

References 306 

17 Determination of Enzyme Activities in Contaminated Soil 309 

Rosa Margesin 

17.1 General Introduction 309 

17.2 Lipase-Esterase Activity 310 

17.3 Fluorescein Diacetate Hydrolytic Activity 313 

17.4 Dehydrogenase Activity 316 

References 319 

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 32 1 

Adolf Eisentraeger, Kerstin Hund-Rinke, Joerg Roembke 

18.1 General Introduction: Strategy 321 

18.2 Sample Preparation 323 

18.3 Water- Extractable Ecotoxicity 330 

18.3.1 Vibrio fischeri Luminescence-Inhibition Assay 330 

18.3.2 Desmodesmus subspicatus Growth- Inhibition Assay.... 331 

18.4 Water-Extractable Genotoxicity 332 

18.4.1 The umu Test 332 

18.4.2 Salmonella/Microsome Assay (Ames Test) 333 

18.5 Habitat Function: 

Soil/Microorganisms, Soil/Soil Fauna, Soil/Higher Plants 334 

18.5.1 Respiration Curve Test 334 

18.5.2 Ammonium Oxidation Test 337 

18.5.3 Combined Earthworm Mortality/Reproduction Test ... 340 

18.5.4 Collembola Reproduction Test 342 

18.5.5 Plant Growth Test 344 

18.5.6 Test Performance for the Derivation 

of Threshold Values 346 

18.6 Combined Performance of Bioassays and Assessment of the 
Results 348 

18.6.1 Water-Extractable Ecotoxic Potential 348 

18.6.2 Water-Extractable Genotoxicity 349 

18.6.3 Assessment of the Habitat Function 350 

18.6.4 Overall Assessment - Combined Strategy 353 

References 355 

Subject Index 361 



Contributors 



Aislabie, Jackie 

Landcare Research, Private Bag 3127, Hamilton, New Zealand 

Braddock, Joan R 

College of Natural Science and Mathematics, University of Alaska Fair- 
banks, Fairbanks, Alaska 99775, USA 

Brookes, Philip C. 

Agriculture and Environment Division, Rothamsted Research, Harpenden, 

Herts., AL5 2JQ, UK 

Douglas, Gregory S. 

NewFields Environmental Forensic Practice LLC, Rockland, Massachusetts 

02370, USA 

Eisentraeger, Adolf 

Institute of Hygiene and Environmental Medicine, Aachen University of 

Technology, Pauwelsstr. 30, 52074 Aachen, Germany 

Foght, Julia 

Biological Sciences, University of Alberta, Edmonton AB, Canada T6G 2E9 

Greer, Charles W. 

Biotechnology Research Institute, National Research Council of Canada, 

6100 Royalmount Ave., Montreal, Quebec, Canada H4P 2R2 

Hedrick, David B. 

Hedrick Services, Knoxville, TN 37932-2575; Center for Biomarker Anal- 
ysis, University of Tennessee, 10515 Research Drive, Suite 300, Knoxville, 
Tennessee 37932-2575, USA 

Hund-Rinke, Kerstin 

Fraunhofer Institute for Molecular Biology and Applied Ecology, P.O. Box 

1260, 57377 Schmallenberg, Germany 



XIV Contributors 

Jarvinen, Olli 

Finnish Environment Institute, P.O. Box 140, 00251 Helsinki, Finland 

Joergensen, Rainer Georg 

Department of Soil Biology and Plant Nutrition, University of Kassel, Nord- 

bahnhofstr. la, 37213 Witzenhausen, Germany 

J0rgensen, Kirsten S. 

Finnish Environment Institute, P.O. Box 140, 00251 Helsinki, Finland 

Klauser, Thomas 

Institute of Microbiology, Leopold Franzens University, Technikerstrasse 

25, 6020 Innsbruck, Austria 

Kucharski, Rafal 

Land Management Department, Institute for Ecology of Industrial Areas, 

Kossutha 6 St, 40-833 Katowice, Poland 

Kuhad, Ramesh C. 

Department of Biotechnology, Kurukshetra University, Kurukshetra - 

136 119, Haryana, India 

Lindstrom, Jon E. 

Shannon & Wilson, Inc., 2355 Hill Road, Fairbanks, Alaska 99709; Institute 

of Arctic Biology, University of Alaska Fairbanks, Fairbanks, Alaska 99775, 

USA 

Malkowski, Eugeniusz 

Department of Plant Physiology, Faculty of Biology and Environmental Pro- 
tection, University of Silesia, Jagielloiiska 28 St, 40-032 Katowice, Poland 

Margesin, Rosa 

Institute of Microbiology, Leopold Franzens University, Technikerstrasse 

25, 6020 Innsbruck, Austria 

Paetz, Andreas 

Deutsches Institut fur Normung (DIN), Normenausschuss Wasserwesen 

(NAW), 10772 Berlin, Germany 

Peacock, Aaron 

Center for Biomarker Analysis, University of Tennessee, 10515 Research 

Drive, Suite 300, Knoxville, Tennessee 37932-2575, USA 



Contributors XV 

Plaza, Grazyna A. 

Institute for Ecology of Industrial Areas, 40-844 Katowice, 6 Kossutha, 

Poland 

Prince, Roger C. 

ExxonMobil Research and Engineering Co., Annandale, New Jersey 08801, 

USA 

Raubuch, Markus 

Department of Soil Biology and Plant Nutrition, University of Kassel, Nord- 

bahnhofstr. la, 37213 Witzenhausen, Germany 

Ripp, Steven 

The University of Tennessee, Knoxville, Tennessee, 37996, USA 

Roembke, Joerg 

ECT Oekotoxikologie GmbH, Boettgerstr. 2-14, 65439 Floersheim, Ger- 
many 

Sainio, Pirjo 

Finnish Environment Institute, P.O. Box 140, 00251 Helsinki, Finland 

Salminen, Jani 

Finnish Environment Institute, P.O. Box 140, 00251 Helsinki, Finland 

Sas-Nowosielska, Aleksandra 

Land Management Department, Institute for Ecology of Industrial Areas, 

Kossutha 6 St, 40-833 Katowice, Poland 

Schinner, Franz 

Institute of Microbiology, Leopold Franzens University, Technikerstrasse 

25, 6020 Innsbruck, Austria 

Singh, Ajay 

Department of Biology, University of Waterloo, Waterloo, Ontario, Canada 

N2L3G1 

Suortti, Anna-Mari 

SGS Inspection Services, Syvasatamantie 24, 49460 Hamina, Finland 

Tien, Albert J. 

Holcim Group Support Ltd Corporate Social Responsibility Occupational 

Health and Safety, Im Schachen, 5113 Holderbank, Switzerland 



XVI Contributors 

Ulfig, Krzysztof 

Institute for Ecology of Industrial Areas, 40-844 Katowice, 6 Kossutha, 

Poland 

Ward, Owen P. 

Department of Biology, University of Waterloo, Waterloo, Ontario, Canada 

N2L3G1 

White, David C. 

Center for Biomarker Analysis, University of Tennessee, 10515 Research 

Drive, Suite 300, Knoxville, Tennessee 37932-2575, USA 

Whyte, Lyle G. 

Dept. of Natural Resource Sciences, McGill University, Macdonald Campus 

21, 111 Lakeshore Road, St. Anne de Bellevue, Quebec, Canada H9X 3V9 

Wilke, Berndt-Michael 

Institute of Ecology, Berlin University of Technology, Franklinstrasse 29, 

10587 Berlin, Germany 



1 



Soil Sampling and Storage 

Andreas Paetz, Berndt-Michael Wilke 



1.1 

Objective of Soil Sampling 
1.1.1 

Principal Objectives 

General 

Samples are collected and examined primarily to determine their physical, 
chemical, biological, and radiological properties. This section outlines the 
more important factors which should be considered when devising a sam- 
pling program for soil and related material. More detailed information is 
given in subsequent sections. 

Whenever a volume of soil is to be characterized, it is generally not pos- 
sible to examine the whole and it is therefore necessary to take samples. 
The samples collected should be as fully representative as possible, and all 
precautions should be taken to ensure that, as far as possible, the samples 
do not undergo any changes in the interval between sampling and exami- 
nation. The sampling of multiphase systems, such as soils containing water 
or other liquids, gases, biological material, radionuclides, or other solids 
not naturally belonging to soil (e.g., waste materials), can present special 
problems. In addition, the determination of some physical soil parameters 
may require so-called undisturbed soil samples for correct execution of the 
relevant measurement. 

Before any sampling program is devised, it is important that the objec- 
tives be first established since they are the major determining factors, e.g., 
the position and density of sampling points, time of sampling, sampling 
procedures, subsequent treatment of samples and analytical requirements. 
The details of a sampling program depend on whether the information 
needed is the average value, the distribution, or the variability of given soil 
parameters. 



Andreas Paetz: Deutsches Institut fur Normung (DIN), Normenausschuss Wasserwesen 
(NAW), 10772 Berlin, Germany 

Berndt-Michael Wilke: Institute of Ecology, Berlin University of Technology, Franklinstrasse 
29, 10587 Berlin, Germany, E-mail: bmwilke@tu-berlin.de 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



2 A. Paetz, B.-M. Wilke 

Some consideration should be given to the degree of detail and precision 
that will be required, and also to the manner in which the results are 
to be expressed and presented, for example, concentrations of chemical 
substances, maximum and minimum values, arithmetic means, median 
values, etc. Additionally, a list of parameters of interest should be compiled 
and the relevant analytical procedures consulted; these will usually give 
guidance on precautions to be observed during sampling and subsequent 
handling of soil samples. 

It may often be necessary to carry out an exploratory sampling-and- 
analysis program before the final objectives can be defined. It is important 
to take into account all relevant data from previous programs at the same 
or similar locations and other information on local conditions. Previous 
personal experience can also be very valuable. Time and money allocated 
to the design of a proper sampling program are usually well justified be- 
cause they ensure that the required information is obtained efficiently and 
economically. 

It is emphasized that complete achievement of objectives of soil inves- 
tigations depends mainly on the design and execution of an appropriate 
sampling program. The four principal objectives of soil sampling may be 
distinguished as follows and are discussed below: 

• Sampling for determination of general soil quality 

• Sampling for characterization purposes in preparation of soil maps 

• Sampling to support legal or regulatory action 

• Sampling as part of a hazard or risk assessmenthack 

The utilization of the soil and site is of varying importance depending on 
the primary objective of an investigation. For example, while consideration 
of past, present, and future site use is particularly relevant to sampling for 
risk assessment, it is less important for soil mapping where the focus is 
on description rather than the evaluation of a soil. Objectives such as soil 
quality assessment, land appraisal, and soil monitoring take utilization into 
account to varying degrees. 

The results obtained from sampling campaigns to assess soil quality for 
mapping may indicate a need for further investigation. For example, if 
contamination is detected, a need arises for identification and assessment 
of potential hazards and risks. 

Sampling for Determination of General Soil Quality 

This is typically carried out at irregular time intervals to determine the 
quality of the soil for a particular purpose, e.g., agriculture. As such, it will 
tend to concentrate on factors such as nutrient status, pH, organic matter 
content, trace element concentrations, and physical factors, which provide 



1 Soil Sampling and Storage 3 

a measure of current quality and which are amenable to manipulation. 
Sampling is usually carried out within the main rooting zone and also 
at greater depths but sometimes without exact distinction of horizons 
or layers. The guidance given in ISO 10381-4 (2003) will be particularly 
relevant. 

Sampling for Preparation of Soil Maps 

Soil maps maybe used in soil description, land appraisal (taxation), and for 
soil monitoring sites to establish the basic information on the genesis and 
distribution of naturally occurring or man-made soils, their chemical, min- 
eralogical, biological composition, and their physical properties at selected 
positions. The preparation of soil maps involves installation of trial pits 
or core sampling with detailed consideration of soil layers and horizons. 
Special strategies are required to preserve samples in their original physical 
and chemical condition. Sampling is nearly always a once-off procedure. 
The guidance given in ISO 10381-4 (2003) is particularly relevant. 

Sampling to Support Legal or Regulatory Action 

Sampling may be required to establish base-line conditions prior to an 
activity that might affect the composition or quality of soil, or it may be re- 
quired following an anthropogenic effect such as the input of an undesirable 
material that may be from a point or a diffuse source. Sampling strategies 
need to be developed on a site-specific basis. To adequately support legal 
or regulatory action particular attention should be paid to all aspects of 
quality assurance including, for example, £C chain-of-custody procedures." 
The guidance given in ISO 10381-5 (1995) is particularly relevant; that in 
ISO 10381-4 (2003) may also be relevant. 

Sampling for Hazard and Risk Assessment 

When land is contaminated with chemicals and other substances poten- 
tially harmful to human health and safety or to the environment, it may 
be necessary to carry out an investigation as a part of a hazard and/or risk 
assessment i.e., to determine the nature and extent of contamination, to 
identify hazards associated with the contamination, to identify potential 
targets and routes of exposure, and to evaluate the risks relating to current 
and future use of the site and neighboring land. A sampling program for 
risk assessment (in this context: phase I, phase II, phase III, and phase IV 
investigations) may have to comply with legal or regulatory requirements, 
and careful attention to sample integrity is recommended. Sampling strate- 
gies should be developed on a site-specific basis. The guidance given in 
ISO 10381-5 (1995) is particularly relevant, and that in ISO 10381-4 (2003) 
may also be relevant. 



4 A. Paetz, B.-M. Wilke 

1.1.2 

Specific Objectives 

General 

Depending on the principal objective(s) it will usually be necessary to 
determine for the body of soil or part thereof: 

• The nature, concentrations, and distribution of naturally occurring sub- 
stances 

• The nature, concentrations, and distribution of contaminants (extrane- 
ous substances) 

• The physical properties and variations 

• The presence and distribution of biological species of interest 

It will often be necessary to take into account changes in the above 
parameters with time, caused by migration, atmospheric conditions, and 
land/soil use. Some detailed objectives are suggested in the clauses below. 
The list is not exhaustive. 

Sampling for the Determination of Chemical Soil Parameters 

There are many reasons for chemical investigation of soil and related ma- 
terial and only a few are mentioned here. It is important that each sampling 
routine is tailored to fit the soil and the situation. Chemical investigations 
are carried out 

1. To identify immediate hazards to human health and safety and to the 
environment 

2. To determine the suitability of a soil for an intended use, e.g., agricultural 
production, residential development 

3. To study the effects of atmospheric pollutants including radioactive fall- 
out on the quality of soil (which may also provide information on wa- 
ter quality and indicate if problems are likely to arise in near-surface 
aquifers) 

4. To assess the effects of direct inputs to soil; there may be contributions 
from: 

- naturally occurring substances that exceed local background values, 
e.g., certain mineral phases in metal deposits 

- (un)expected contamination by application of agrochemicals 

- (un)expected contamination due to industrial processes 



1 Soil Sampling and Storage 5 

5. To assess the effect of the accumulation and release of substances by soils 
on other soil horizons or on other environmental compartments, e.g., 
the transfer of substances from a soil into a plant 

6. To study the effect of waste disposal, including the disposal of sewage 
sludge on a soil (which, apart from contributing to the pollution load, 
may produce other chemical reactions such as the formation of persistent 
compounds, metabolites, or the evolution of gases, such as methane) 

7. To identify and quantify products released by industrial processes and by 
accident (usually done by investigation of suspect sites or contaminated 
sites) 

8. To evaluate soil derived from construction works with view to possible 
or further utilization of such soils or disposal as waste 

Commonly, sampling strategies are employed that require samples to 
be taken either from identifiable soil horizons or from specified depths 
(below ground surface). It is best to avoid mixing the two approaches, 
particularly when sampling natural strata, as this can make it difficult to 
compare results. However, a coherent combination of the two approaches 
can sometimes be useful on old industrial sites where there is variation in 
both the nature of fill and the depth of penetration of mobile contaminants 
into the ground, i.e., where there are two independent reasons for changes 
in soil/fill properties. 

Knowledge of the way in which particular chemical substances tend to be 
distributed between the different compartments (air, soil, water, sediment, 
and living organisms) is advantageous for the design of some sampling 
programs. Similarly, knowledge of the behavior of living organisms affected 
by chemical substances, or that influence the availability of substances due 
to microbiological procedures, is also advantageous. 



Sampling for the Determination of Physical Soil Parameters 

The sampling of soil for the determination of some physical properties re- 
quires special consideration since the accuracy and extrapolation of mea- 
sured data rely on obtaining a sample that retains its in situ structural 
characteristics. In many circumstances, it may be preferable to conduct 
measurements in the field since the removal of even an undisturbed sample 
can change the continuity and characteristics of soil physical properties and 
lead to erroneous results. However, certain measurements are not possible 
in the field. Others require specific field conditions, but the field situation 
can only be controlled to a very limited extent; e.g., it may be possible to 
modify the hydrological situation temporarily for measurement purposes 
by irrigation. The time and expense necessary for field measurements may 



6 A. Paetz, B.-M. Wilke 

not be affordable. Laboratory measurements of physical properties are 
therefore frequently necessary. 

Differences and changes in soil structure affect the choice of size of 
sample. Hence, a representative volume or minimum number of replicates 
must be determined for each soil type to be studied. The moisture status of 
the soil at sampling can influence physical measurements, e.g., hysteresis 
on rewetting can occur. Many physical properties have vertical and lat- 
eral components, and this should be considered prior to sampling. Where 
small undisturbed soil samples are required, manual excavation of cores, 
clods, or soil aggregates can be applied. Sampling equipment should be de- 
signed such that minimal physical disturbance to the soil occurs. For larger 
samples, the use of hydraulic sampling equipment and cutting devices is 
preferable in order to obtain a sample with minimal disturbance. Care 
should be taken in both equipment design and manufacture to ensure that 
internal compression or compaction of the sample does not occur. Where it 
is difficult to obtain an undisturbed sample for laboratory measurements, 
e.g., in stony or iron pan soils, then in situ measurements may be the most 
appropriate method. 

Sampling for the Assessment of Biological Soil Parameters 

Biological soil investigations address a number of different questions re- 
lated to what is happening to or caused by life forms in and on the soil, 
including both fauna and flora in the micro and macro range. Ecotoxico- 
logical questions are usually given first priority. For example, tests should 
be made to verify the effects of chemicals added to the soil on life-forms 
and also the possible effects of life-forms in the soil on plants (e.g., high- 
value crops) and on the environment, especially on human health. In some 
cases, biological soil test procedures operate with fully artificial soils, but 
normally the major task of sampling is to choose a reliable soil or site to 
carry out the tests. Sampling for the assessment of aerobic microbial pro- 
cesses is covered in ISO 10381-6 (1993). The sampling for the assessment 
of anaerobic processes is described in ISO 15473 (2002). 

1.2 

Selection of Sampling Technique 

The selection of appropriate sampling equipment depends on the objective 
of sampling and should be done after consideration by the analyst or 
scientist responsible for subsequent determination. ISO 10381-2 (2002) 
gives guidance on commonly used equipment for sampling soil and related 
material. Parts 4, 5, and 6 of ISO 10381 describe needs for specific purposes 
within their scopes. 



1 Soil Sampling and Storage 7 

1.3 

Sampling Strategy 

1.3.1 
General 

The strategy for the site investigation (whether preliminary, exploratory, 
or main) will be determined by the objectives. For example, the different 
requirements of site investigations for the purpose of selling, determining 
whether contamination is present as suspected, or redevelopment will influ- 
ence the spacing of sample locations and the number of samples analyzed, 
and hence the cost of the investigation. 

Before embarking on any phase or stage of investigation it is important 
to set data quality objectives in terms of the type, quantity, and quality (e.g., 
analytical quality) of the data and other information to be collected. These 
data quality objectives will depend in part on the nature of the decisions 
to be made on the basis of the investigation and the confidence required in 
those decisions. Failure to set data quality objectives at the outset can lead 
to considerable waste of money if, for example, the data collected are not 
suitable or sufficient for a reliable hazard assessment, or leave too many 
uncertainties about the "conceptual model" developed for the site. 

1.3.2 

Preliminary Investigation 

General 

This is an investigation comprising a desk study (see below) and site recon- 
naissance (walk-over survey, site inspection). It is carried out using histori- 
cal records and other sources to obtain information on the past and present 
usage of the site together with information about local soil properties, ge- 
ology, hydrogeology, and environmental setting. From this investigation, 
the possibility of contamination can be deduced, and hypotheses can be 
formulated on the nature, location, and distribution of the contamination. 
These hypotheses form part of the overall conceptual model of the site 
that should be developed, encompassing not only the contamination as- 
pects but also the geology, hydrogeology, geotechnical properties, and en- 
vironmental setting. The current and planned site uses are also important 
aspects of the conceptual model. The preliminary investigation should 
provide sufficient information: 

• For initial conclusions about potential hazards and hazards to actual or 
potential human and other receptors, and 

• For determination as to need for further action. 



8 A. Paetz, B.-M. Wilke 

The amount and type of information required will depend on the objec- 
tives of the investigation and the ease with which the information can be 
obtained, i.e., the amount of work required will vary with the age of the 
site, the complexity of its historic usage, the complexity of the underlying 
geology, etc. 

It shall be remembered that the contamination on a site may be more 
complex than initially indicated (for example by current usage) and ade- 
quate information on site history should always be obtained in the prelim- 
inary investigation. 

Desk Study 

This includes collection of relevant information on the site, e.g., location, 
infrastructure, utilization, history. Possible sources of this information are 
publications, maps (check accuracy of map used), aerial photographs, and 
satellite imagery from, e.g., land surveyor's offices, geological surveys, water 
management boards, industrial inspection boards, mining boards, mining 
companies, geotechnical institutions, regional and local (city) archives, 
agricultural and forestry authorities, and building supervisory boards. 
Particularly important is information on the physical and chemical prop- 
erties and the possible spatial distribution of the soil parameter under 
investigation; special attention must be paid to geological features such as 
stratigraphy and hydrogeology. 

Site Reconnaissance 

A visit of the site should be part of the preliminary investigation, prefer- 
ably in conjunction with the desk study, although it may be independent. 
Depending on the local variability of the site and the technical difficulty 
of the planned investigation, an experienced person should be chosen for 
this task. Such a visit gives a first impression about the correlation of ex- 
isting maps with reality, and it will provide much additional information 
in a comparatively short time. In some cases, it may be necessary to draw 
a first or additional map at this stage. 

Samples are not often taken during preliminary investigations; if they 
are, they are usually needed to obtain an overview of the kind of soil in 
order to chose the right equipment for later activities. Parts 4, 5, and 6 of 
ISO 10381 specify the range of preliminary investigations used within their 
scopes. 

Output from Preliminary Investigation 

A report should be prepared summarizing the findings of the preliminary 
investigations and stating the conclusions (or hypotheses) drawn concern- 



1 Soil Sampling and Storage 9 

ing the anticipated site conditions (e.g., geology, hydrology, possible con- 
tamination) relevant to the design of the sampling program. This should 
enable the appropriateness of the sampling strategy adopted to be assessed 
at a later date. 



1.3.3 

Exploratory Investigation 

This involves on-site investigation including collecting samples of soil or 
fill, surface water, groundwater, and soil gas, where appropriate, to be 
analyzed or tested. The data and information produced are then assessed to 
determine if the hypotheses from the preliminary investigation are correct 
and, where appropriate, to test other aspects of the conceptual model. It is 
therefore mainly a qualitative investigation rather than quantitative. 

In some cases, where the hypotheses are found to be correct, no further 
investigation may need to be carried out. However, it may become apparent 
as a result of the exploratory site investigation, for example, that the con- 
tamination pattern is more complex or concentrations of contamination 
are higher than anticipated and may have already caused or in the future 
may cause a hazard. In this situation the information obtained may be 
inadequate to make decisions with a satisfactory degree of confidence. It 
will be necessary to carry out a main site investigation to produce sufficient 
information to make a full hazard assessment, to determine the need for 
protective or remedial measures, and in due course (and possibly following 
further stages of investigation), to select, design, and apply protective or 
remedial measures. 



1.3.4 

Main Site Investigation 

The main site investigation quantitatively determines the amount and spa- 
tial distribution of contaminants, their mobile and mobilizable fractions, 
and the possibilities of spread into the environment. Also included is the 
possible future development of the contamination situation. This will in- 
volve the collection and analysis of soil or fill, surface water, ground water, 
and soil gas samples in order to obtain the information necessary to enable 
a full assessment of the hazards presented by the contamination to humans 
and other potential receptors and also to enable appropriate containment 
or remediation actions to be identified (sometimes) together with an initial 
estimate of costs. The analysis of samples can be supported by model cal- 
culations and investigation techniques that do not make use of sampling. 



10 A. Paetz, B.-M. Wilke 

Detailed design of protective or remedial works may require further stages 
of investigation. 

The amount and nature of the information required from the main site 
investigation (or any particular stage of it) will vary depending on the 
nature of the site and the objectives of the investigation. The implications 
of the decisions on what actions should be implemented on a site will vary 
from site to site. Additionally, the amount and quality of the information 
required will also vary according to the requirements of the decision- 
making processes (e.g., the risk assessment, decisions regarding the need 
for and type of remedial actions). All parties involved in the decision- 
making process should be kept fully informed as information is produced 
to ensure that the information is sufficient for the purpose intended. 

After completion of the interpretation of the information generated, 
including any risk assessment, it should be possible to determine whether 
protective or remedial measures are required and to make generalizations 
about the type of measures that might be appropriate. 

1.3.5 

Samples and Sampling Points 

General 

The selection, location and preparation of the sampling points depend on: 

• The objectives of the investigation 

• The preliminary information available 

• The on-site conditions 

The nature of samples to be obtained shall be appropriate to the aim of 
the investigation and shall be specified in the program before fieldwork 
begins. 

Sampling Patterns 

Sampling patterns are based on the estimation of the distribution of soil 
constituents (in most cases chemical substances) in an area or, when appro- 
priate, on the type of substance input. Four major fixed sampling patterns 
can be identified as being based on: 

• No specific estimate of substance distribution 

• Local substance distribution and known as a "hot spot" 

• Distributions along a line 

• Strip-like distributions 



1 Soil Sampling and Storage 1 1 

Along with these, several other patterns exist (e.g., based on deposition 
of substances from the air, input due to flooding). All fixed patterns have 
to be adjusted to local conditions and are subject to modification. 

In agricultural sampling a small number of convenient sampling patterns 
are established in order to obtain information on, e.g., nutrient demand 
or pesticide residues of rather large areas. For additional information refer 
to ISO 10381-4 (2003). However, it must be emphasized that most grid 
sampling patterns are not very efficient during the growing season, and are 
rarely applicable. The investigation of contaminated sites which may have 
profound health and economic consequences usually requires a much more 
detailed selection and application of sampling patterns, to give calculated, 
estimated, or randomly chosen sampling points on a one-, two- or three- 
dimensional figure. The choice of pattern should be the result of preliminary 
investigation of a site rather than of an ad hoc decision taken in the field. 

Some investigations are carried out without predetermined pattern 
plans. This should not be confused with the application of random dis- 
tribution of sampling points, because a person usually cannot distribute 
sampling points randomly without preparation, i.e., without ensuring that 
at every point in the area, despite the position of the other sampling points, 
a sample will be obtained with equal probability. Where sampling is to be 
carried out without a predetermined pattern (ad hoc sampling) care shall 
be taken that sampling is carried out by an appropriately experienced in- 
vestigator. It also should not be confused with the application of sampling 
plans to verify special hypotheses which, with regard to the problem, will 
be developed and justified by the investigator (judgmental sampling). 

In the following are examples of a number of commonly applied sampling 
patterns which meet different statistical requirements (Figs. 1.1-1.5). Expe- 
rience (and theoretical considerations) show that in many cases systematic 
sampling on a regular grid is both practical and sufficiently productive 
to allow the creation of a detailed picture of variations in soil properties. 
The number of sampling points can be easily increased (e.g., in areas mer- 
iting more detailed investigation), the grid is easy to mark out on site, 
and sampling points are usually easily relocated. Systematic sampling can 
be supplemented by judgmental sampling when appropriate. ISO 10381-5 
(1995) provides examples of pattern application for sampling contaminated 
sites. For selection of sampling patterns see Fig. 1.5. 

Most natural properties of the soil vary continuously in space and, as 
a consequence, the values at sites that are close together are more similar 
than those further apart. They depend upon one another in a statistical 
sense. This property is known as spatial dependence and its implications 
for sampling are covered by methods of geostatistics, i.e., spatial statis- 
tics. Viewed mathematically, the value of a soil property at any place is 
a function of its position. The only practicable approach is to regard such 



12 A. Paetz, B.-M. Wilke 

a property as a random variable and to treat its variation in space statisti- 
cally. Such properties are known as regionalized variables. The application 
of regionalized variable theory by developing variograms is a common tool 
in geostatistics. 

Another geostatistical approach is multi-stage or nested sampling and 
analysis which also can be linked with a regionalized variable theory. 

The applicability of geostatistical methods does not depend on the ob- 
served values at those sites, but on the configuration of the sampling points 
in relation to the area (or block if three dimensions are considered) to be 
estimated. A general criterion for the usefulness of a sampling pattern lies 
in the smallness of the largest parcel not being sampled. In terms of sta- 
tistically efficient sampling, a regular equilateral triangular grid provides 
the best selection of sampling points. For a grid with one node per unit 
area, the sampling points are 1.0746 units of distance apart, and no other 
point is more then 0.6204 units of distance away from a sampling point. For 
practical purposes, sampling patterns are based on rectangular grids. For 
such a grid with one node per unit area, no point is more than 0.7071 units 
of distance from a sampling point, i.e., the greater ease of use of the square 
grid is offset by the slightly greater area of unsampled site. 



Example 1: Non-Systematic Patterns (Irregular Sampling) 

Widely used in agricultural/horticultural land investigations are the "N", 
"S", " W", and "X" patterns of sampling (Fig. 1.1). The general premise is that 
the distribution of soil constituents is relatively homogeneous. The patterns 
used are simplifications of the stratified random sampling method. Along 
the outline of such a pattern, a number of samples are taken and then may 
be bulked and mixed to provide one sample for analysis. The distribution 
of sampling points is likely to be inadequate to provide the location of point 
pollution, and in any event high contaminant levels will be lost in mixing 
of these samples. Thus, in most contaminated-land investigations, these 
patterns are unlikely to be useful because they obscure high levels of point 
contamination. Wherever there are likely to be differences in soil type or 
conditions, crop growth, plant species, previous cultivation, etc., the site 
should be subdivided according to these differences and a separate sample 
taken from each area. 

Sampling along a single diagonal of a field or a unit is only recom- 
mended in case of strip-like distribution of contaminants on agricultural 
areas due to application of fertilizers. Applying a diagonal for sampling 
avoids systematic bias by simple and effective means, which would arise 
with strip-parallel sampling. However, the more diagonals, the better. Two 
diagonals (X-shape) introduce a serious bias to the central area of the field 
(Fig. 1.1). This should be considered for the evaluation of the results of the 



1 Soil Sampling and Storage 



13 




Fig. 1.1. Non- systematic patterns 

determinations. Application of diagonal patterns should be based on the 
following: 

• Estimation that substances are distributed uniformly 

• Recognition of usefulness only for uniformly developed areas, and of the 
need to sample deviating parts separately 

• Application of more than one diagonal if possible (e.g., parallel or X- 
shape) 

• Equidistant placing of sampling points for all diagonals, i.e. shorter 
diagonals have fewer sampling points 

• Selection of sampling point independent of local characteristics, points 
being fixed (preferably by pacing) 

Traversing the area in a zigzag manner similar to that shown in Fig. 1.2 
is another way of applying a non-systematic pattern. 




Fig. 1.2. Zigzag traverse 
sampling pattern 



14 



A. Paetz, B.-M. Wilke 



A general exception to the "biased diagonals pattern" was developed for 
permanently monitored areas within selected sites to achieve information 
about long-term changes due to human influence. The aim is to make sam- 
ples available from an area representative of the surrounding environment 
for a defined number of examinations to be carried out over a period of 
some years. The following procedure is recommended (Fig. 1.3): 

1. Select a representative area of approx. 1,000 m 2 . 

2. Divide this area into four squares, each of 250 m 2 . 

3. Within each square draw two diagonals, along each of which nine samples 
are obtained (Fig. 1.3). 

4. Take samples according to the specified requirements. 

5. Prepare composite samples 1, 2, and 3 by: 

- Mixing single samples of positions 1, 4, 7, 10, 13, and 16 to give 
composite sample 1 

- Mixing single samples of positions 2, 5, 8, 11, 14, and 17 to give 
composite sample 2 

- Mixing single samples of positions 3, 6, 9, 12, 15, and 18 to give 
composite sample 3 




Fig. 1.3. Rotating diagonals 
pattern for permanently 
monitored areas 



1 Soil Sampling and Storage 



15 



6. Rotational sampling of the area may be conducted by: 

- Taking samples in the intersections of the sampling points (positions 
1-18 in Fig. 1.3) 

- Rotating the diagonals clockwise around the center of the square in 
steps of 22. 5° so that, all in all, four series of samplings can be carried 
out at undisturbed positions 

An area selected and sampled according to the above-mentioned scheme 
serves for eight sampling series. After the final series, the area may be 
considered unsuitable for further sampling. Extensions or reductions in 
the dimensions of the test area may impose changes in the total number of 
samples, and thus also affect composite samples. 

Example 2: Circular Grids 

Circular grids are useful for delineating local contaminations, such as from 
storage tanks, but also for indicating influence around a regional emitting 
source, e.g., precipitation from an industrial plant. Sampling is carried out 
at the intersection of concentric circles (the radii of which will depend on 
the suspected area of contamination) and the radial lines of the main eight 
points of the compass (Fig. 1.4). Sampling based on circular grids may lead 
to information on: 

• Substance concentrations at the grid center (maximum values) 

• Distribution of contamination (size of particular area with increased 
contamination) 

• Shape of distribution of contamination 




Fig. 1.4. Circular grid 



16 A. Paetz, B.-M. Wilke 

Disadvantages of circular grids are: 

• Star-shaped (radial) location of sampling points is practicable but not 
optimal. Rotation of concentric circles by 22. 5° leads to a higher quality 
pattern (Fig. 1.3). 

• Relationship of sampling-point densities of the (usually) eight samples 
close to the center and those (usually) eight samples at greater distance 
might not to be optimal in every case. If, for example, borders of distri- 
bution of a contaminated area are looked for, fewer central points should 
be sampled and more toward the margins of the grid. 

• Circular grids might imply a uniform extension of contamination in 
all directions. This is usually not the case. Preferred directions, e.g., 
due to main wind direction in case of airborne contaminants, should be 
considered in modifying of the circular grid, e.g., an increased number of 
sampling points in critical directions, an extended distance of sampling 
in critical directions. 

• Circular grids generally do not serve for taking composite samples be- 
cause the values thus measured give information neither on the average 
nor on the maximum concentration in the area sampled. 



Example 3: Systematic Sampling (Regular Grids) 

In many cases a regular grid is selected (Fig. 1.5). Because there is a direct 
relationship between optimal sampling point distance and the (estimated) 
dimension of the contamination, spacing between sampling points should 
not exceed the greatest (estimated) extent of the contamination. Grid di- 
mensions will depend on how much detail is required. The assigned spacing 
will differ according to the objective of sampling, e.g., to collect samples of 
average degree of contamination, to locate isolated sources of contamina- 
tion, or to establish the extent of contaminated zones (horizontal and verti- 
cal). The latter is of particular importance in cases where a contamination 
is already located and a follow-up sampling program becomes necessary. 
Although more frequently used for the investigation of soil contamination, 
regular grids are also suitable for soil fertility investigations, general soil 
monitoring programs, etc. An advantage of a regular grid is that it may 
be set up easily and grid dimensions may be readily varied. Interpolation 
between sampling points and return to the grid to carry out a more in- 
tensive sampling in localized areas to further delineate point sources of 
contamination is easy. It is also possible to fix the sampling points at the 
intersections of the grid lines. 



1 Soil Sampling and Storage 



17 



x 



' 


• 


/• JM 


# A 


• 


• 


• 




• / 






/V y I^v. 


// 


• 




• 




y / /2^>L 


/*%// 




/ • 




• 


• 


• 






• 



Fig. 1.5. Regular distribution of sampling points on a regular grid hatched areas indicate 
contamination 



Example 4: Random Sampling 

In cases of presumed irregular occurrences of contaminated zones, random 
sampling may be applied. Sampling points within the area are selected by 
using random numbers, which can be found in tables included in manuals 
on statistics or which may be generated by computer programs. This tech- 
nique has the disadvantage of irregular coverage and makes interpolation 
between sampling points difficult (Fig. 1.6). In general, random sampling 
can also be applied for soil fertility investigations, etc. In practice, random 
sampling (in its purest form) is rarely used in soil surveys. 



Example 5: Stratified Random Sampling 

This method avoids some of the disadvantages of strictly random sam- 
pling. The site is divided into a number of grid cells, and a given number 
of randomly distributed sampling points is chosen in each cell (Fig. 1.7). 
In general, stratified random sampling can also be applied for soil fertility 
investigations, etc. The method has disadvantages in terms of interpola- 
tion between the sampling points. Further sampling of the site to identify 
local areas of contamination based on the original sampling locations is 
difficult. 



18 



A. Paetz, B.-M. Wilke 



x 




Fig. 1.6. Random sampling without grid 
x 






• 


W^W./ 


• 
# ^ 


y% 


• 


• 


/• 




w/%\ / 




fyf // 


^^ • 


• \ 


mm 


I / n^>L 


J* jMz 1 


/wfyfrJ^ 


/ • 


• 


• 


•^ 


^<4m iffi 




• 



Fig. 1.7. Stratified random sampling 



1 Soil Sampling and Storage 



19 



Example 6: Unaligned Random Sampling 

The term "unaligned" means "irregular" in the sense of "not-in-a-line." 
The method is similar to stratified random sampling but in this case only 
one of two coordinates is chosen at random. The procedure is as follows. 
For example: given a grid with 24 cells (squares), arranged in 4 lines and 
6 columns (Fig. 1.8): 

1. For the first cell (line 1, column 1), x- and y-coordinates are chosen at 
random. 

2. For cells 2, 3, 4, 5, and 6 only the y-coordinates are chosen at random. 

3. For cells 7, 13, and 19, only the x-coordinates are chosen at random. 

4. All sampling points are now located on the grid: For all sampling points 
in the columns, the y-coordinates of cells 2, 3, 4, 5, and 6 are valid, and 
for all sampling points in the lines the x-coordinates of cells 7, 13, and 
19 are valid. 

The method has disadvantages in terms of interpolation between the 
sampling points. Further sampling of the site to identify local areas of 
contamination based on the original sampling location is difficult. 



x 




Fig. 1.8. Unaligned random sampling on a regular grid 



20 



A. Paetz, B.-M. Wilke 



Example 7: Systematic Sampling on a Non-Rectangular Grid 

In case of an equilateral triangular grid (Fig. 1.9), each grid point is neigh- 
bored by three grid points at the unique distance d x . No other adjacent 
points exist. The free, unsampled distance between the related adjacent 
points has a radius of 



3 



(1.1) 



The circular area (A) not being sampled therefore is 



d 2 
A = n • r ' = n • — 

3 



(1.2) 



Example: given an area of 10 x 10 m and using 99 sampling points arranged 
in 11 rows with 9 sampling points each (distance between rows = 1.11 m) 
the area not being sampled is 1.29 m 2 . This unsampled area is thus smaller 
than for example a rectangular grid of the same size and using 100 sampling 
points arranged at a distance of 1 m one from another, where the area not 
being sampled is 1.57m 2 . Any circular contamination with r > 0.64m is 
certain to be detected. Thus, just by changing the pattern (and with one 
sample less) the size of the unsampled area decreases to approx. 18%. 

Sampling points at the site are fixed at a distance of d x in parallel rows 
spaced at a distance 



d y — — • V3 



(1.3) 



That is, approx. 0. 87 x d x . The sampling points on the parallel rows are 
staggered by 



X 

~2 



(1.4) 




Fig. 1.9. Triangular grid 



1 Soil Sampling and Storage 



21 





• 


m 




a 


m 


41 




A 


• 




\ 




W 
/ 




w 




V 




V 








V 

X 








■ 




• 








• 




V 

2x 












• 








• 









Fig. 1.10. Sampling along 
a linear source 



Example 8: Sampling Along a Linear Source 

In case of contamination following a line, e.g., caused by leaking pipelines, 
sampling points can be arranged in the covering soil directly above the 
pipeline or, if not practicable for certain reasons, close to the pipeline. If 
the distribution of contaminants caused by a line-like structure is also of 
interest, it is recommended to take samples at a distance x one from another 
above the line and further samples at increasing distances (e.g., 2x) parallel 
to the line (Fig. 1.10). 



Identifying the Sampling Location 

Identification of sampling points is not usually necessary when taking 
composite samples for agricultural purposes. Where samples are taken at 
pre-defined points, their accurate location and identification is important 
for three principal reasons: 

1. To enable actual sampling locations to be revisited if necessary 

2. To enable accurate plotting of data in relation to site features so that any 
needed treatment (e.g., additions of nutrients or removal of contamina- 
tion) can be properly planned 

3. To enable the data to be stored and processed by computers (e.g., for 
modeling studies, preparation of maps, input into geographic informa- 
tion systems) 

Moreover, it is recommended that a sketch map be prepared present- 
ing all relevant information on the sampling location. Both maps and 
photographs should include a scale and a direction marker. It is impor- 
tant for the interpretation of data, particularly on abandoned industrial 
sites to have detailed information on surface levels at sampling locations. 



22 A. Paetz, B.-M. Wilke 

Sampling locations should be determined with an appropriate degree of 
accuracy. Because it may be necessary to vary the actual location away from 
the predetermined location because of the presence of obstructions, it may 
be preferable to do the accurate surveying of sampling locations once the 
sampling exercise is completed or as it progresses. Surface levels can be 
determined at the same time. 

When investigating abandoned industrial, waste disposal, or other po- 
tentially contaminated sites, the horizontal and vertical location of sam- 
pling points or probing points should be recorded. The location of sampling 
points should be marked before sampling begins using poles/markers with 
color sprays. Color sprays should not be used if soil air has to be sampled. 

Preparation of the Sampling Site 

Depending on the objective of the investigation, a sampling pattern is 
chosen at the design stage and is then applied in the field. Within the 
range of patterns are some very complex ones developed with the help of 
computer-aided statistics. Preparation for sampling with the use of such 
patterns, e.g., location of desired sampling points on the ground, can be 
very time-consuming, especially when samples are to be obtained by bor- 
ing/drilling techniques or from trial pits. Preparation of the site includes, 
for example, removal of superficial deposits (e.g., uncontrolled deposition 
of urban wastes), establishment of safety measures, installation of mea- 
surement devices (if field tests are carried out together with sampling), as 
well as exactly locating the sampling points. In many cases, preparation 
of the site takes longer than the actual sampling procedures. Both during 
and on completion of sampling all necessary measures must be taken to 
avoid hazards to the health and safety of anyone entering the site, and to 
the environment. 

Barriers to Sampling 

It may not be possible to sample at a planned location due to a variety 
of reasons (e.g., trees, large rocks, buildings, buried foundations or public 
utility services, difficulties of access) and contingency plans for dealing with 
such situations should be made in advance. The action to take will depend 
on the circumstances. The investigator may ignore the unavailable point 
or follow predetermined rules for choosing a nearby substitute location 
(e.g., alternative position within 10% of grid spacing or paired sampling 
along grid lines on either side the obstruction). Ad hoc decisions made in 
the field can lead to bias. An attempt should be made when mapping out 
the site to identify such obstructions in advance of actual field work. In all 
cases when a sampling point has to be relocated, this fact, and the reasons 
for relocation, should be clearly indicated in the report. 



1 Soil Sampling and Storage 23 

Preliminary investigations as described in Sect. 1.3.2 should provide as 
much detail as possible about conditions expected to exist on the site and 
should therefore guide the design and execution of the sampling program. 
However, such investigations cannot totally prevent the danger of misin- 
terpretation of the results of borings, and the selection of sampling points 
should take this into account. 

Depth of Sampling 

No general recommendation can be given on the depths at which samples 
should be taken or on the final depths to which trial pits or boring/ drilling 
should extend. This depends on the objectives and might be subject to 
change during a running program. Investigation of soil for chemical char- 
acteristics can be divided into two general types: 

1. The investigation of agricultural and similar near-natural sites, where 
information is required mostly on the topsoil or plowed horizon or arable 
zone but often over an extended area. 

2. The investigation of sites which are known or suspected to be contami- 
nated, where information is required from deeper layers, sometimes to 
a depth of several tens of meters, the extent of the area usually being 
rather small compared to agricultural sites. 

A mixture of both cases is realized in so-called "soil-monitoring sites," 
which represent larger areas of homogeneous soil development and in most 
cases are established to monitor environmental effects to the complete 
profile over a long-term scale. A precise description should be made of 
all soil horizons or layers encountered during the sampling exercise and 
included in the report. 

If a profile is to be sampled, care should be taken that every horizon/layer 
of interest is sampled and that different horizons/layers are not mixed. In 
general, contaminated sites should be sampled horizon by horizon unless 
stated otherwise by the client. Care should be taken in a site investigation 
to ensure that pathways for migration of contamination are not created, 
particularly where impermeable strata maybe penetrated. 

When trial pits are used it may be appropriate to sample from more 
than one site. A depth-related sampling program is based on a number 
of conventions, depending on the project. It is not as representative with 
regard to the soil as a horizon-related sampling program can be. The 
mode of sampling from each depth should be carefully specified; e.g., the 
maximum depth range (usually not more than 0.1 m) and how horizontal 
variations are to be dealt with. 

The total depth reached, the thickness of the horizons/layers penetrated, 
and the depth from which the samples are obtained should be recorded. All 



24 A. Paetz, B.-M. Wilke 

data should be recorded in meters below surface. The soil depth should be 
measured from the ground surface with the thickness of the humus litter 
layer recorded separately. 

Mountain regions or hilly areas with pronounced slopes require special 
consideration. For slopes of 10° and greater, vertical drilling lengths should 
be extended according to the cosine rule in order to maintain constant 
slope-parallel thicknesses of soil layers. The extension factor is 1/cos of 
slope. Without correction, for example, the error will be 2% at a slope of 
11.5°. 

Timing of Investigation 

In some circumstances, it may be necessary to restrict sampling to specific 
periods of the year. For example, if the characteristic or substance to be 
determined is likely to be affected by seasonal factors or human activities 
(weather, soil conditioning/fertilization, use of plant protecting agents), this 
should be taken into account in the design of the sampling program. This is 
particularly important where monitoring continues for several months or 
years or is repeated periodically, and therefore requires similar conditions 
every time sampling is carried out. 

Sample Quantity 

At least 1,000 g of fine soil should be obtained for chemical analysis. This 
figure applies both to single samples and composite samples, in the latter 
case after sufficient homogenization. Samples obtained to serve as reference 
material or to be stored in a soil specimen bank should be of larger size, 
usually larger than 2,000 g. 

Where the sampling of soil involves the separation of oversized material 
(i.e., mineral grains, sand, pebbles, and all other materials) due to very 
coarse-grained or heterogeneous soil conditions, the material removed 
shall be weighed or estimated and recorded and described to enable the 
analytical results to be given with reference to the composition of the 
original sample. These procedures should be carried out in accordance 
with ISO 11464(1994). 

Details on the amount of sample materials needed for determination 
of specific physical soil parameters are given in the respective methods 
(Chap. 2). In particular, the determination of the particle size distribution 
may need a very large mass of soil material. The actual mass required will 
usually depend on the largest grain size to be determined (see ISO 11277 
1998). The quantity of soil sample needed for biological or ecotoxicological 
investigations is highly dependent on the aim of the investigation and the 
related soil organisms. 



1 Soil Sampling and Storage 25 

Single Samples vs. Composite Samples 

Composite samples are usually required in cases where the average con- 
centration of a substance in a defined horizon/layer is to be determined. 
Single samples are required in cases in which the distribution of a sub- 
stance over a defined area and/or depth is sought. In most guidelines on 
sampling for agricultural or similar investigations, it is recommended that 
composite samples be collected by taking a number of increments (accord- 
ing to ISO 10381-4 (2003) at least 25 increments should be obtained) and 
combining them to form a composite sample. When preparing composite 
samples regard should be paid to analytical requirements. For example, 
composite samples should never be used if volatile compounds are to be 
determined. 



1.4 

Sampling Methods 

1.4.1 
General 

The most commonly used methods of sampling and forming holes in the 
ground to collect samples are covered in this text. This does not preclude 
the use of other techniques that are suited to the problems of a partic- 
ular location, e.g., areas of permafrost, nor does it preclude the use of 
other methods that have been developed. Whatever technique is used, the 
principles of sample collection and the approach to sampling to obtain an 
appropriately representative sample should be adhered to. This will include 
the minimization of contamination of the sample and the protection of the 
samplers and other personnel involved. The choice of sampling method 
will be determined by taking into account all the needs of the investigation, 
including distribution of sampling locations, size and type of sample, and 
the nature of the site, including any problems the site poses in carrying out 
the investigation. 

1.4.2 

Type of Sample 

There are three basic approaches to taking samples from the ground for 
the purpose of investigating soil and ground conditions. A sample maybe: 

Type 1 Material collected from a single point (disturbed or undisturbed 
sample). 



26 A. Paetz, B.-M. Wilke 

Type 2 A composite of small incremental point samples taken close to- 
gether [disturbed sample; perhaps not suitable for certain tests, 
e.g., the determination of volatile organic compounds; (VOCs)]. 

Type 3 A composite of small incremental point samples taken over an area 
(such as a field; disturbed sample). 

Samples taken to identify the distribution and concentration of particu- 
lar elements or compounds will normally be samples of type 1 or perhaps 
type 2 within the area being examined. Such samples would be appropriate 
for geological or contamination investigations and any other investigation 
involving disturbed samples. Samples taken to assess the overall quality or 
nature of the ground in an area would be type 3. Such samples would be 
taken for agricultural purposes. 

Disturbed samples may be taken by any of the three basic methods since 
these samples do not require the maintenance of the original ground struc- 
ture. Undisturbed samples will always require type 1 sampling because the 
original ground structure needs to be retained in the sample. Undisturbed 
samples can be taken using a coring tool or cylinder or with a sampling 
frame. Whichever of these sampling devices is used, the mode of operation 
is the same. The sampling device is pushed into the ground to be sam- 
pled and then subsequently removed complete with the sample so that the 
ground is collected in its original physical form. 

Type 1 samples can be readily collected using hand augers and other 
similar sampling techniques. Any of the following tools (as well as others) 
may be used as appropriate: 

• Cutting cylinders of different size, cutting frame 

• Special hand augers [gauge auger (shallow-profile sampler), bucket auger 
to bring down borings for cutting cylinder application]; 

• Protective cap, hydraulic or handpowered supporting ring 

Special bags should be used for storage and transport of "sample rings" 
(actually sample cylinders of limited height) to prevent disturbance and 
drying out. Where undisturbed samples are required, special equipment 
(see above) will be necessary in order to collect the sample while maintain- 
ing the original ground structure. 

Type 2 samples will be appropriate when using machines for excavating 
ground to obtain samples. In these circumstances the samples should be 
formed by taking portions from locations within the bucket of excavated 
material (e.g., nine-point sample, according to Fig. 1.4). 

Type 3 samples can be collected using hand or powered augers, but care 
needs to be taken to ensure the auger repetitively collects the same amount 
of sample. 



1 Soil Sampling and Storage 27 

Disturbed samples are suitable for most purposes except for some physi- 
cal measurements, profiles, and microbiological examinations when undis- 
turbed samples maybe required. Undisturbed samples should be collected 
where it is intended to determine the presence and concentration of VOCs, 
since disturbance will result in loss of these compounds to the atmosphere. 

Choices of sampling method include the use of machinery or manual 
methods. The sampling may be carried out near the ground surface, at 
some depth below ground level, or from locations deep below the ground 
surface. Methods of achieving the desired depth for sampling are either by 
excavating (e.g., trial pits), driving probes, or drilling (e.g., boreholes). 

Sampling during borehole creation allows the required integrity for the 
chemical, physical, and biological investigation of selected soil horizons. 
Gas and water sampling may also be undertaken for specific purposes re- 
lating to the need to acquire information rapidly, for example monitoring 
a borehole for methane and carbon dioxide or VOCs on occasions when the 
rapid identification of chemical constituents in groundwater is required. It 
is recommended that monitoring groundwater horizons over time for hy- 
drogeological and chemical parameters, as well as ground composition, be 
undertaken from cased wells or standpipes installed in boreholes. The re- 
quirements of the sampling strategy should identify the nature of borehole 
construction so that the appropriate monitoring design can be specified. 

1.4.3 

Undisturbed Samples 

If undisturbed samples are required for soil sampling, these can easily 
be taken, for example, using a Kubiena box, a coring tool, or cylinder. In 
each case the sampling device is pushed into the soil and subsequently 
removed with the sample so that the soil is collected in its original physical 
form. Beside these simple techniques, many others exist, some of which are 
described later. 

Hand-Operated Auger Techniques 

There are many designs of hand auger samplers available. The designs 
have been developed over many years to deal with different soil types 
and conditions. Ease of use depends upon the nature of the ground to be 
sampled. In general, handaugers are easier to use in a sandy soil than in 
other soils, particularly where obstructions such as stones are encountered. 
In sandy soils, hand augers can be used to sample to a depth of about 
5 m. Hand augers are usually used for sampling homogeneous soils, e.g., 
agricultural soils. When using hand augers, care should be taken to ensure 
that the soil is not contaminated by material dropping into the sample from 



28 A. Paetz, B.-M. Wilke 

higher up the bore either during augering or during withdrawal of the 
samples. Lining the borehole carefully with a plastic tube can prevent this 
cross contamination. 

Preferred forms of hand augers to be used for collection of soil samples 
are those which take a core sample. Other types of auger may be used to 
facilitate drilling to the requisite depth for sampling providing it is possible 
to clean the bore to prevent cross contamination. 

Sampling by hand augers allows observation of the ground profile and 
the collection of samples at preselected depths. Particular care should 
be taken to obtain representative samples if localized contamination is 
penetrated. When a hand auger is to be used to take samples for testing 
soil for agricultural purposes, and the samples are to be composited, it 
is essential that the auger should be capable of consistently collecting the 
same sample volume. Such sampling of the near-surface soil is normally 
done at approx. 150-250 mm depth. 

Power-Operated Auger Techniques 

It is possible to obtain augers powered by small motors to reduce the labor 
required to carry out the sampling. The need to avoid cross contamination 
within the bore applies equally to augering with power-operated augers 
as with hand augers. Powered augers mounted on rough-terrain vehicles 
are available for repetitive sampling for agricultural purposes. Care should 
be exercised when using fuel-driven motors to avoid contamination of the 
sample by the fuel, the motor lubricant, and the exhaust fumes. Augers 
powered by electric motors that minimize the risk of such contamination 
are available. 

Light Cable Percussion Boring 

Light cable percussion boring general uses a mobile rig with winch of 1-2 1 
capacity driven by a diesel engine and a tripod derrick of about 6 m height. 
With many types the derrick folds down so that the rig can be towed by 
a small vehicle (frequently four-wheel drive). The light cable percussion 
technique is commonly used for geotechnical purposes, and boreholes 
over 20 m deep can be created. This technique can be of particular use in 
investigating deep sites such as refuse tips and other unstable ground. The 
ground is penetrated using different tools, depending on the strata. A clay 
cutter is used for cohesive soils and a shell (or bailer) for cohesionless 
soils. Chisels may be used to penetrate very hard ground and obstructions. 
The borehole formed by these tools is supported by a steel casing that is 
advanced as the borehole proceeds. 

Depending upon the nature of the ground, the tool may form the borehole 
in advance of the steel casing being pushed down the hole, e.g., in clay 



1 Soil Sampling and Storage 29 

strata. This often results in material from the side of the borehole being 
dislodged as the casing is pushed down the borehole, and can result in 
cross-contamination. If the borehole is being formed in sands or gravels, 
particularly in the saturated zone, the steel casing may be pushed into place 
to support the borehole sides before the material is removed with the shell. 
This can disturb the ground and make sampling difficult. 

In some strata it may be necessary to add water to the borehole to 
provide lubrication. In this situation tap water may be used, if available, 
and care should be taken with respect to the effects on both soil and 
water samples. The addition of water should be recorded on the borehole 
log and, if appropriate, on the sample details. The clay cutter and the 
shell bring up disturbed material from the borehole which is generally 
sufficiently representative to permit recording of the strata, but care has 
to be taken to avoid misinterpretation due to ground being pushed down 
within the borehole - for example, when the casing is moved. The casing 
avoids most of the problems of cross contamination, but the borehole 
should be cleaned out each time the supporting casing is driven further 
into the borehole, before taking a sample. Samples may be collected from 
both the clay cutter and the shell. The resultant sample size, although larger 
than obtained by hand-augering techniques, is still restricted. Undisturbed 
samples may be collected in cohesive strata and in weak rock (e.g., chalk) 
by driving a hollow tube (100 mm open-tube sampler) into the ground and 
withdrawing the resultant core for examination and analysis. Use of such 
undisturbed sampling equipment may be preferred in order to minimize 
cross-contamination of samples collected for testing purposes. 

Water samples may be obtained as drilling proceeds and, because the 
casing of the borehole seals the borehole from the surrounding ground as 
the borehole advances, it is possible to sample water horizons at different 
depths with minimal risk of cross contamination. However water samples 
that are truly representative of the ground water necessitate the installation 
of an appropriately designed monitoring well. The borehole atmosphere 
can be monitored for gas concentrations as the borehole proceeds, or gas 
samples maybe taken so that the profile of the ground gas composition can 
be determined. 



Rotary Drilling 

Powered rotary cutting tools use a shaft fitted with a cutter head that 
is driven into the ground as it rotates. The system requires some form 
of lubrication (air, water, or drilling mud) to keep the cutting head cool 
and remove the soil and other material that has been cut through. The 
lubricant lifts the debris from the cutting head up the borehole formed 
and ejects the material at ground level. This results in the potential for 



30 A. Paetz, B.-M. Wilke 

cross contamination due to contact with the ground forming the sides of 
the hole. This technique is particularly useful for digging a hole quickly 
in order to form a deep observation well or for obtaining samples using 
a technique appropriate at greater depths only. The uncontrolled ejection 
of material that can occur with this technique (for instance where air or 
water is used for lubrication) can lead to extensive surface contamination 
when drilling through contaminated ground. This may be hazardous, both 
to the investigation team and the environment. 

There are two basic types of rotary drilling, (1) open hole (or full hole) 
drilling in which the drill cuts all the material within the diameter of the 
borehole, and (2) core drilling where an annular bit fixed to the bottom 
of the outer rotating tube of the core barrel assembly cuts a core that is 
recovered within the inner most tube of the core barrel assembly and is 
brought to the surface for examination and testing. Rotary drilling requires 
well-maintained equipment operated by a specialist driller with adequate 
training and considerable experience. 

Driven Auger 

The driven auger is powered by machine, so that great force can be exerted 
downwards. The cutter head consists of one or more 360° spirals, usually 
with a shallow pitch to prevent ground falling off when withdrawn from 
the borehole. The method of forming the borehole is to advance the cutter 
head approx. 1 m into the ground, withdraw the head from the hole and spin 
off the spoil. This process is repeated until the required depth is reached. 
This method is not very satisfactory for sampling, because of the potential 
for cross contamination, nor is it suitable for strata logging. The method 
does enable the formation of a large diameter hole (up to 25 cm) into the 
ground relatively quickly. Lubrication of the auger is not required, but some 
dispersal of contaminated material may occur as the spoil is spun from the 
cutter head. 

Continuous Flight Auger 

A similar system is the continuous flight auger, which consists of a contin- 
uous helix welded to the center shaft. Downward force is again provided by 
the machine and continuous rotation lifts the ground to the surface from 
the base of the hole. This technique is only of use in site investigations in 
forming a hole rapidly to give depth in the ground and cannot be used for 
sampling or strata logging. Lubrication of the auger is not required. 

Hollow Stem Auger 

Hollow stem augers are a form of continuous flight auger in which the 
continuous helix is attached to a hollow central shaft. The drill head is 



1 Soil Sampling and Storage 31 

formed of two pieces, a circular outer head and an inner pilot or center bit 
that is fixed on a plug on the hollow shaft that can be withdrawn through 
the center of the auger up to the surface. This ability to withdraw the center 
bit and plug whilst leaving the auger in place is the principal advantage of 
the hollow stem auger. Withdrawing the plug provides an open cored hole 
into which samplers, undisturbed samplers, instruments, borehole casing, 
and numerous other items can be inserted to the depth achieved. 

Removal of any such equipment and replacing the center plug and bit 
enables the continuation of the borehole. The technique provides a fully 
cased hole and can avoid some of the potential cross-contamination prob- 
lems of percussion boring. Ground samples are collected by open drive 
samplers or core barrels inserted down the hollow stem. The method has 
been successful on some landfill sites and can be used for the installation 
of groundwater monitoring wells and gas standpipes. Some versions of the 
hollow stem auger allow continuous access to the bottom of the borehole 
and will permit percussion drilling or driven sampling through the center, 
while the hollow stem auger is actually forming the hole. The technique will 
allow collection of samples, particularly undisturbed samples, in addition 
to other down-hole testing, and also enables strata logs to be produced. 
Lubrication of the auger is not required. 

Percussive window sampling involves driving cylindrical steel tubes into 
the ground using a high frequency percussive hammer. Usually, the hammer 
is driven by a hydraulic power pack, but electric and pneumatic hammers 
are also available to suit particular site conditions. Sample tubes are 1 or 2 m 
long and have a broad slot or window cut down one side. The soil material 
passes into the sample tube, through a cutting shoe at the end, as it is driven 
into the ground. Drill rods are used to drive the sample tubes to greater 
depths. On reaching the required depth for sampling, the sample tube and 
any drill rods are withdrawn using a mechanical jack. After removal from 
the probe hole, the soil material can then be inspected and the strata logged 
and sampled from the window. 

Soil samples may also be obtained using split tubes or split spoon sam- 
plers. These are effectively tubes linearly split in half but held together by 
securing rings during sampling. Such devices are often used in conjunc- 
tion with driven bar probes, and they allow ready retrieval of the core. Soil 
samples may also be obtained using a tube combined with an inert liner 
to enable ease of removal of the core from the sampler. The system can be 
used to collect samples at different depths, to rapidly penetrate to the depth 
at which the sample is to be taken, or to provide a continuous core. 

Sample tubes of various diameters are available (35-80 mm) and se- 
lected according to the ground conditions. Tubes are normally selected in 
a sequence of reducing diameters to penetrate to depth. The depth that 
can be achieved depends on the soil type and particularly on the presence 



32 A. Paetz, B.-M. Wilke 

(or absence) of obstructions. Depths of 10-12 m can be achieved where 
the probe hole remains open without support. Piezometers and ground gas 
monitoring pipes can be installed in the resultant probe holes where the 
ground is sufficiently stable. Systems are available to allow a probe head, 
with a sampling device, to be inserted into the previously formed hole to the 
desired sampling depth. The probe head is then unscrewed and withdrawn 
up the inside of the shaft, and the exposed sampling device is pushed into 
the ground to collect the sample. The sampling head is then withdrawn and 
removed for analysis. This system also enables undisturbed samples to be 
collected. 

Continuous Samplers 

Continuous soil samplers can produce core samples up to 30 m length in 
ground such as fine alluvial deposits. This may be of particular value and 
is considered to yield superior samples to those obtained by consecutive 
drive-in sampling. The samplers normally are made in sizes between 30 
and 70 mm diameter and consist of an outer driven tube with an internal 
system providing a sheath to the core as the sampler is driven into the 
ground. Extension tubes of 1 m length are added to the sampler as the 
ground is penetrated. On removal from the ground, the continuous core is 
cut to suitable lengths, frequently 1 m, and placed in purpose-made sample 
cases for storage. Samples may be removed from the core for testing and 
the core itself observed and recorded. 

Driven Probes 

Driven probes maybe used to make continuous geophysical measurements, 
for example, resistance to penetration, or may be fitted with instruments 
for gathering other data. Care should be taken to avoid cross contamination 
from the sides of the probe hole and from the base of the probe hole. This 
system can be used to either monitor ground water parameters (such pH, 
electrical conductivity, temperature, etc.) using monitors in the probe, or to 
access groundwater so that a representative sample can be taken without the 
need for purging as associated with conventional monitoring wells. Ground 
gases can be similarly accessed and sampled. Driven probes have the usual 
disadvantage of difficulty in penetrating ground with obstructions, and 
cannot be used for logging the ground strata unless continuous soil samples 
are taken. Driven probes are, however, considerably faster than traditional 
boreholing techniques. 

Excavations (Trial Pits) 

This is a widely used technique for collecting samples for site investigations 
related to contamination. The advantages of the method are the applicability 



1 Soil Sampling and Storage 33 

over a wide range of ground conditions, the opportunity for close visual 
examination of the strata, and the speed with which the work can be 
carried out. Trial pits can be dug where the ground will stand temporarily 
unsupported and permit the observation of the in-situ condition of the 
ground both vertically and laterally. Where there is water present in the 
excavation, problems are presented due to instability of the sides and the 
difficulty of obtaining representative samples of the ground (finer material 
tends to wash out with the water as the sample is collected). In this situation 
the trial pit may be dewatered by pumping, providing there is a safe and 
suitable means of disposal of the water - or an alternative technique of 
sampling should be used. In deeper trial pits formed by machines, samples 
of the ground can be collected by careful use of the machine bucket, thereby 
avoiding any need to enter the pit. In carrying out excavations, whatever 
technique is used to form a trial pit, the excavated material should be 
placed on the adjacent ground (this should be protected as necessary from 
contamination) in a way that ensures it will not fall back into the excavation 
causing cross contamination. 

The surface soil layer should be kept separate so that it can be replaced 
on the surface after the trial pit is backfilled. It may be necessary to separate 
other material as it is excavated so that any deep lying contamination is 
replaced at the same depth when back filling and not mixed with other 
material or replaced near the surface. For environmental reasons and due 
to legislation, it may be necessary to dispose of excavated material off-site 
and to complete the backfilling of the trial pit and restoration of the site 
using clean imported material. 

Entry of the excavation by personnel should be avoided where possible 
since the unsupported sides of a trial pit may readily collapse. If it is 
essential that an excavation is to be entered for sampling purposes, e.g., 
the collection of undisturbed samples, then shoring should be used and 
reference should be made to the guidance given in ISO 10381-3 (2001). In 
unstable ground the trial pit may collapse and extra care should be taken 
when observing the excavation and collecting samples. If necessary, the 
sides should be supported or made to slope to improve stability. For all 
ground conditions, if the depth of excavation is greater than 1-1.2 m and 
the excavation is to be entered by personnel, the sides should be adequately 
shored to prevent collapse. 



Manual 

Shovel, pick, and fork may be used to excavate trial pits down to about 
2 m and, if only a small number of such excavations are required, this may 
be the easiest technique for collecting soil samples. The trial pit should 
have a plan area of approx. 1 x 1 m to enable easy collection of samples 



34 A. Paetz, B.-M. Wilke 

and recording of the soil profile. Hand excavation is necessary particularly 
in urban areas if services (water, gas, electricity, etc.) are known to exist 
in the vicinity, and particularly if their location is uncertain. Once the 
base of the excavation is below the depth at which any services may exist, 
then the excavation or boreholing may be continued using the appropriate 
machinery. 

1.4.4 
Cross-Contamination 

Whatever method is used, it is important that nothing connected with the 
sampling system itself contaminates the sample. This includes avoiding 
contamination by contact with the sampling equipment or containers and 
also avoiding the loss of contaminants from the sample by adsorption or 
volatilization. The sampling equipment should be kept clean so that parts 
of a previous sample are not transmitted to a subsequent sample causing 
cross contamination. For agricultural purposes, even with repetitive sam- 
pling across a field to form a composite sample, the sampling device should 
at least be brushed clean between each location. For geological and con- 
tamination investigations, all sampling equipment should be thoroughly 
cleaned between each sample. Contamination of samples due to lubrica- 
tion used to ease sample collection, or contamination due to equipment 
lubricants, oils, greases, or fuels should be avoided. Where it is necessary 
to use lubrication, e.g., water, to ease forming a borehole to enable sample 
collection, only lubrication that will not conflict with nor confound the 
analysis of the samples (in the sense of matrix effects or contribution to the 
contamination) should be used. 

A hand trowel of stainless steel should be used to place samples into 
sample containers. The quality of the stainless steel should, however, first 
be verified to ensure that cross contamination of the samples will not occur 
or interfere with the quality of the analytical data. The most commonly 
used methods of drilling, excavating, and sampling of the ground produce 
disturbed samples. If undisturbed samples are required, special sampling 
equipment is required and extra care should be taken in collection. 

1.4.5 

Sampling Containers 

General Considerations 

Samples of soils and related materials are liable to change to differing 
extents as a result of physical, chemical or biological reactions that may take 
place between the time of sampling and the analysis. This is especially true 



1 Soil Sampling and Storage 35 

of soils contaminated with volatile constituents. The causes of variations 
are numerous and may include: 

• Changes of certain constituents due to the activities of living organisms 
in the soil 

• Oxidation of certain compounds by atmospheric oxygen 

• Changes in the chemical nature of certain substances due to changes of 
temperature, pressure, and hygroscopicity (e.g., loss to the vapor phase); 

• Modification of pH, conductivity, carbon dioxide content, etc., by the 
absorption of carbon dioxide from the air 

• Irreversible adsorption on the surface of containers by metals in solution 
or in a colloidal state, or by certain organic compounds 

• Polymerization or depolymerization 

The extent of these reactions is a function of the chemical and biological 
nature of the sample, its temperature, its exposure to light, the nature 
of the container in which it is placed, the time between sampling and 
analysis, conditions such as rest or agitation during transport, seasonal 
conditions, etc. It must be emphasized, moreover, that these variations are 
often sufficiently rapid so as to modify the sample considerably within 
several hours. It is therefore essential in all cases to take the necessary 
precautions to minimize these reactions, and in the case of many parameters 
to analyze the sample with a minimum of delay. Any of the procedures 
should be mentioned in the sampling report if applied during sampling. 

Preservation 

The addition of chemical preservatives or stabilizing agents is not a com- 
mon practice for soil sampling. This is because a single soil sample is usually 
used for a large number of different determinations, and moreover has to 
undergo preparation (drying, milling, etc.) during which unwanted and 
unquantifiable reactions of the preservatives may occur. If, in special cases, 
it is necessary to preserve samples a method that does not introduce un- 
acceptable contamination should be chosen. Generally, stability of samples 
can be considered in three classes: 

1. Samples in which the contaminant(s) is/are stable 

2. Samples in which the contaminant(s) is/are unstable but stability can be 
achieved by a preservation method 

3. Samples in which the contaminant(s) is/are unstable and cannot be 
readily stabilized 



36 A. Paetz, B.-M. Wilke 

For those contaminants that are unstable, loss or change (chemical or 
biological) of the contaminant should be minimized by either preserving 
the contaminant (e.g., freezing or adding a stabilizing agent) or by arrang- 
ing for analysis to be undertaken immediately or soon after sampling. The 
use of liquid nitrogen for immediate deep freezing of soil samples in vapor 
phase is effective, and containers made of stainless steel (not chromium or 
nickel plated) are recommended. Some contaminants are not easily stabi- 
lized in a manner compatible with subsequent analysis. Volatile solvents 
fall into this category and some of them may begin to volatilize as soon as 
the soil is exposed by sampling. A special sampling procedure is needed 
to minimize such loss. In spite of numerous investigations carried out in 
search of methods that will enable soil samples to be stored without modifi- 
cation of their composition, it is impossible to give absolute rules that cover 
all cases and all situations and that do not have exceptions. In every case, 
the method of storage must be compatible with the analytical techniques 
to be used and should be discussed with the analytical laboratory. 

Use of Appropriate Containers 

The choice and the preparation of containers can be of major importance. 
The most frequently encountered problems are: 

• Adsorption onto the walls of the containers 

• Improper cleaning resulting in contamination of the container prior to 
sampling 

• Contamination of the sample by the material of which the container is 
made 

• Reaction between constituents of the sample and the container 

The purpose of the container is to protect the sample from losses due to 
adsorption or volatilization, or from contamination by foreign substances. 
Other factors to be considered in selection of the sample container used to 
collect and store the sample include: 

• Resistance to temperature extremes 

• Resistance to breakage 

• Water and gas tightness 

• Ease of reopening 

• Size, shape, and mass 

• Availability 

• Potential for cleaning and re-use 



1 Soil Sampling and Storage 37 

Cleaning of the sample container is a very important part of any sam- 
pling/analysis program. Two basic situations can be distinguished: ( 1 ) clean- 
ing of new containers to remove dust and packing material; (2) cleaning of 
used containers prior to re-use. The type of cleaners used depends on the 
kind of container material and on the material to be analyzed. The selection 
of acids or other cleaning agents should ensure that no contamination of 
the containers results with regard to the constituents to be analyzed and, 
moreover, that there is no harm to the environment or human health. 

Containers already used for investigations of contaminated sites should 
not be used again because cleaning containers of soils containing unknown 
substances may cause risks to health. The determination of organic con- 
stituents may require drying or cooling procedures under carefully con- 
trolled conditions to avoid microbial contamination. Sterilization is re- 
quired whenever biological or microbiological determinations are to be 
carried out. 



1.5 
Pretreatment 

1.5.1 

Chemical Analysis 

Inorganic Parameters and Soil Characteristics 

Soil samples are dried in the air or in an oven at temperature not exceeding 
40 °C, or are freeze-dried. If necessary, the soil sample is crushed while still 
damp and friable and again after drying. The soil is sieved and the fraction 
smaller than 2 mm is divided into portions mechanically or by hand, to 
enable representative subsampling for analysis. If small subsamples (< 2 g) 
are required for analysis, the size of the particles of the fraction smaller 
than 2 mm is further decreased. 

• A drying temperature of 40 °C in an oven is preferable to air drying 
at room temperature because the increased speed of the drying limits 
changes due to microbial activity. 

• It should be noted that every type of pretreatment will have an influence 
on several soil properties. 

• The sieve aperture size of 2 mm is generally used. However, before the 
pretreatment is started, check should be made to see if any of the ana- 
lytical methods to be applied require other sieve sizes. 

• Storing soil samples, including samples that are air dried, refrigerated 
or stored in the absence of light, for a long time may have an influence 



38 A. Paetz, B.-M. Wilke 

on a number of soil parameters, especially solubilities of both inorganic 
and organic fractions. 

• Special measures should usually be taken for samples from contaminated 
soils. It is important to avoid contact with the skin, and special measures 
should be taken when drying such samples (ventilation, air removal, 
etc.). Samples may be hazardous because of the presence of chemical 
contaminants, fungal spores, or pathogens such as leptospirosis, and 
appropriate safety precautions should be taken. 

• According to the international standard, it is generally assumed that at 
least 500 g of fresh soil shall be available. 

• Keeping an archive sample is optional and should be clearly stated in the 
overall description of the investigation program. 

Organic Contaminants 

The properties of organic micro-pollutants may differ greatly according to 
chemical species: 

• They can range from non volatile to very volatile compounds (low to 
high vapor pressure). 

• They may be labile or reactive at ambient or elevated temperatures. 

• They may be biodegradable or UV degradable. 

• They may have considerably different solubilities in water. 

• They require different analytical procedures. 

Because of these differences a general pretreatment procedure cannot be 
proposed. The goal of a pretreatment procedure is to prepare a test sample 
in which the concentration of the contaminant is equal to the concentration 
in the original soil, provided, however, that this procedure does not alter the 
chemical species to be analyzed. If the sample contains only small particles 
and the contaminant is homogeneously distributed it is, for instance, not 
necessary to grind the sample. According to the international standard the 
size 2 mm is used to distinguish between small and large soil particles. Care 
should be taken to ensure consistency among the following aspects: 

• Soil diversity 

• The aim and accuracy of the analysis 

• The nature of the chemical species to be analyzed 

Important to pretreatment is the particle size distribution of the sample 
in relation to the mass of sample taken for analysis. For the analysis of 



1 Soil Sampling and Storage 39 

organic contaminants, the mass taken in most cases is about 20 g. With 
such a sample mass, and provided that the contaminant is homogeneously 
distributed and the particles in the sample are smaller than about 2 mm, 
further grinding of the sample is not necessary. If the sample contains large 
particles or if the contaminant is heterogeneously distributed (for instance, 
tar particles), it is not possible to take a representative test sample of about 
20 g without grinding the sample. To improve the homogeneity, samples 
are grinded to a size smaller than 1 mm. Prior to analysis very often no 
information about the distribution of the contaminant in the soil is known. 
Some analytical procedures start with a field-moist sample. Drying of the 
sample will give lower extraction results, But because the sample is not dry, 
grinding is not possible. In a situation in which accurate results are needed, 
the best available pretreatment procedure should be used. If it is necessary 
to establish whether the concentration is above a certain limit, and it is 
already known that the soil is heavily polluted, the simplest pretreatment 
procedure will perhaps meet the needs despite drawbacks. In that case, 
however, the result may have to be presented as not representative of the 
whole sample. 

Three methods for the pretreatment of soil samples in the laboratory 
prior to the determination of organic contaminants are applied in routine 
analysis: 

1. A method for pretreatment if VOCs are to be measured. Core test sam- 
ples are taken from the sample and extracted according to the specific 
analytical procedure. If composite samples are required, extracts of indi- 
vidual samples are mixed. It is usually not possible to obtain composite 
samples without severe losses of volatiles. 

2. A method for pretreatment of moderately volatile to non-volatile organic 
compounds where the result of analysis must be accurate and repro- 
ducible. The sample contains particles larger than 2 mm and/or the con- 
taminant is heterogeneously distributed: Samples are chemically dried 
at a low temperature (-196 °C, liquid nitrogen). The freeze-dried sam- 
ples are ground with a cross beater mill with a sieve of 1 mm (cryogenic 
crushing). After grinding suitable test portions are processed according 
to the specific analytical procedures. Composite samples can be prepared 
by mixing of the ground samples. If the extraction procedure prescribes 
a field-moist sample, drying and grinding is not possible. If the original 
samples only contain a small fraction of particles greater than 2 mm and 
the distribution of contaminants is likely to be homogeneous, grinding 
may be omitted. In these two cases suitable test portions are directly 
taken after mixing of the sample. To distinguish more volatile from less 
volatile organic compounds, boiling points are used instead of vapor 
pressure at ambient temperature. For some specific components in the 



40 A. Paetz, B.-M. Wilke 

group of moderately volatile compounds, freeze drying may give good 
results. (In the International Standard freeze drying is not described.) 

3. A method for pretreatment if non volatile organic compounds are to be 
measured and the extraction procedure prescribes a field-moist sample, 
or if the largest particles of the sample are smaller than 2 mm and the 
contaminant is homogeneously distributed, mixing by hand is the only 
pretreatment that need be applied. This procedure may also be used if 
reduced accuracy and repeatability are acceptable. 

The choice depends above all on the volatility of the organic compounds 
under analysis. It also depends on the soil particle size distribution, the 
heterogeneity of the sample, and the analytical procedure that is to follow. 

1.5.2 

Physical Analysis 

Usually, the determination of soil physical parameters requires undisturbed 
soil samples. Thus, pretreatment plays only a minor role. Exceptions are: 

• Determination of the water content, which can be carried out to support 
calculation of the analytical result, i.e., to standardize the result on dry 
soil mass. In this case, the analysis can be performed in the laboratory 
based on disturbed soil sample material. On the other hand, if the soil 
water content needs to be determined on a volume basis, an undisturbed 
sample must be taken and no further treatment applied before testing. 

• The determination of the particle size distribution. Depending on the 
range of particle sizes, the nature (chemistry) of the soil material, and the 
objective of the investigation, a suite of different pretreatment procedures 
maybe applied, including drying, slightly breaking aggregates, removing 
specific kinds of materials, chemically breaking aggregates down, etc. 
The matter is very complex and needs specialists' advice in most cases. 

1.5.3 

Biological Analysis 

As a rule, soil for microbiological analyses under laboratory conditions 
should be sampled in the field with a water content that facilitates sieving. 
In the laboratory the soil should be processed (sieving) as soon as possible 
after sampling. Soil fauna and plant tests can be also carried out on air-dried 
samples. In this case the samples must be pretreated in order to achieve 
optimum conditions for the species present. 



1 Soil Sampling and Storage 41 

1.6 

Storage of Samples 

1.6.1 
General 

• Soils samples for laboratory determinations are collected in many stud- 
ies. In general the samples are taken at the site, mixed, or otherwise 
treated at the site, packed in containers, and then transported to the lab- 
oratory. Upon arrival at the laboratory the samples may again be treated 
before being sent for analysis. Some samples may be stored directly for 
later analysis. After analysis the samples may be discarded or stored. 
The samples are stored when there is a need for further analysis, either 
because parameters already determined require rechecking or a need 
exists for making additional determinations in the future. 

• The conditions for storage should be selected carefully at all stages from 
the point of taking the sample. Examples of storage conditions are light, 
temperature, humidity, accessibility, duration of storage, type of con- 
tainers, and amount of storage. The documentation is also important. 
Risk and security problems should be considered. Well-designed storage 
conditions, such as provisions for monitoring, are particularly impor- 
tant in large-scale studies where the number of samples may become 
quite large over the years. Incorrectly chosen storage conditions may 
lead to high costs and may render the samples unfit for future use. 

• The effect of storage on biodiversity is not discussed because of the 
difficulty to define this parameter. 

• Radioactivity decay is generally not affected by storage and is not treated 
in this standard. Radioactive change caused by loss or gain of matter 
should be considered in connection with the appropriate compounds. 

• Containers holding samples should be protected and sealed in such 
a way that the samples do not deteriorate or lose any part of their content 
during transport. Packaging should protect the containers from possible 
external contamination, particularly near the opening, and should not 
itself be a source of contamination. Most of the analytical procedures 
used in chemical soil analysis recommend that soil samples be taken to 
the laboratory immediately after sampling, but in some cases a range of 
time is given during which the sample should arrive in the laboratory. 

• Soil samples should be kept cool and dark during transportation and 
storage. 



42 A. Paetz, B.-M. Wilke 

• Cooling or freezing procedures can be applied to increase the period 
available for transport and storage. A cooling temperature of 4 ± 2°C 
has been found suitable for many applications. But cooling and freezing 
procedures should only be used in consultation with the analytical lab- 
oratory. Freezing especially requires detailed control of the freezing and 
thawing process in order to return the sample to its initial equilibrium 
after thawing. 

• Light-sensitive soil constituents require storage in darkness or, at least, 
in light-absorbent containers. 

• Undisturbed samples should be transported in the absence vibration or 
other physical disorder in order to maintain the original structure. 

• Disturbed samples, and especially non-cohesive, very dry soils, tend to 
separate into different particle fractions during transportation. In such 
cases the soil material should be re-homogenized before pretreatment 
and analysis. 

• Any national regulations regarding the packaging and transport of haz- 
ardous materials should be observed. 



1.6.2 

Specific Considerations for Biological Parameters 

Biological tests can be separated into soil microbiological, fauna, plant, and 
biodegradation tests, and tests for the ecotoxicological characterization of 
soils and soil materials. Storage conditions for soils used for these tests vary 
over a wide range and depend on the organism or parameter to be tested. 

Microbiological Tests 

Samples should be stored in the dark at 4 ± 2 °C with free access of air. It 
is preferable to use soils as soon as possible after sampling. If storage is 
unavoidable, this should not exceed 3 months unless evidence showing con- 
tinued microbial activity is provided. The active soil microflora decreases 
with storage time, even at low temperatures, and the rate of decrease de- 
pends on the composition of the soil and the microflora involved (see also 
ISO 10381-6 1993). 

If soil samples have to be stored for longer periods than 3 months, 
freezing of samples at -20, -80, or -150 °C may be appropriate, although 
not generally recommended. It has been shown for a number of soils from 
temperate climates that storage at -20 °C for up to 12 months does not 
inhibit microbial activity (e.g., ammonium oxidation). Soil samples for 
phospholipid fatty acid (PLFA) and DNA analyses can be stored at -20 °C 



1 Soil Sampling and Storage 43 

for 1-2 years. Samples for rRNA analyses can be stored at -80 °C for the 
same period. In the latter case the samples should be frozen immediately 
at -180 °C (shock freezing with liquid nitrogen). 

Longer storage periods are mainly needed if the influence of added 
pollutants on soil microbes and microbial processes has to be tested with 
the same soil material, or if the community structure (structural diversity; 
PLFA, DNA, RNA) of soils has to be evaluated at a distinct point of time 
during the year. In these cases the time needed for analyses can easily 
exceed 3 months (chemical, pollutant testing). For structural analyses of 
the microflora, storage at -4 °C is not suitable. 

If longer storage of samples at temperatures below -20 °C is used, special 
attention has to be given to the thawing of samples. Freeze-thaw cycles can 
increase the availability of organic matter to micro-organisms (Haynes and 
Beare 1996). For analyses of microbial activity (e.g., soil respiration) a thaw- 
ing period of 1 day at 4°C and another 3 days at 20 °C is recommended. 
Generally drying of soils is not recommended although air drying and 
rewetting is a common physiological stress for the microbial communities 
in surface soils. It has been shown that drying- rewetting events can induce 
significant changes in microbial carbon and nitrogen dynamics that can 
last for more than a month after the last stress (Fierer and Schimel 2002). 
Rewetting after drying causes bursts of respiration and growth of distinct 
populations of bacteria (Lund and Goks0r 1980). 

Investigations indicate that fast thawing (1 day at 20 °C in an incubator) 
results in smaller variations in microbiological parameters (e.g., microbial 
activity) than the recommended slow thawing when compared to a frozen 
control (Weinfurtner et al. 2002). 

Biodegradation Tests 

For testing the biodegradation of organic chemicals in soils (ISO 11266 
1994; ISO 15473 2002), storage of soils should be avoided if possible because 
activity of soil microorganisms will decrease in the course of time. Storage 
at 4 °C up to 3 months is permissible. For the assessment of degradation 
of chemicals in anaerobic soils under anaerobic conditions, the access of 
oxygen should be avoided during storage. 

Tests Involving Soil Fauna and Higher Plants 

There are no specific recommendations for soil storage with respect to 
soil fauna and higher plant tests in ISO standards. It is recommended to 
store the soil samples under the same conditions as for testing of microbes 
and microbial processes. The reason for this is that the availability and 
effectiveness of pollutants is essentially governed by microbial activity. 
The same is also true for plant testing. Additionally, the nutrient supply of 



44 A. Paetz, B.-M. Wilke 

test soils should be considered, especially if unknown contaminated soils 
are tested, to avoid false negative results. 

Ecotoxicological Testing 

Generally, sieved samples should be stored in darkness. For microbial 
analyses, soils and soil materials should be handled as described above. 
For terrestrial analyses (e.g., plant tests, earthworm tests) samples can be 
stored at 4 ± 2 °C for 3 months. For testing the leaching potential/ retention 
function of soils and soil materials, water extracts for aquatic tests should 
be prepared immediately after sieving. If the tests cannot be performed 
within 7 days (storage of the extracts at 4 ± 2°C in the dark), extracts 
should be stored at -20 °C. 



1.6.3 

Preparing the Samples After Storage 

The procedures for preparing the samples after storage will depend on the 
storage conditions and the analyses. It is not possible to give a general spec- 
ification. Existing standards (e.g., ISO 11464 1994) have to be considered. 
When a soil sample is stored for a long period of time, a vertical redistri- 
bution may occur. A new mixing in a suitable mixer is advisable. For large 
samples, this may not be sufficient. It is recommended that the sample 
be spread in a thin layer on a plastic foil, and then the layer repeatedly 
folded and spread it out again. Especially, the conditions of thawing have 
to be defined because this can influence the determination of biological, 
microbiological, and organic parameters. The soil samples stored below 
°C must be unfrozen in original bags or containers. 

References 

Fierer N, Schimel P (2002) Effects of drying- rewetting frequency on soil carbon and nitrogen 
transformations. Soil Biol Biochem 34:777-787 

Haynes RJ, Beare MH (1996) Aggregation and organic matter storage in meso-thermal, 
humid soils. In: Carter MR, Steward BA (eds) Soil structure and organic matter storage. 
CRC/Lewis, Boca Raton, pp 213-262 

ISO 10381-1 (2002) Soil quality - Sampling - Part 1: Guidance on the design of sampling 
programmes 

ISO 10381-2 (2002) Soil quality - Sampling - Part 2: Guidance on sampling techniques 

ISO 10381-3 (2001) Soil quality - Sampling - Part 3: Guidance on safety 

ISO 10381-4 (2003) Soil quality - Sampling - Part 4: Guidance on the procedure for investi- 
gation of natural, near- natural and cultivated sites 

ISO 10381-5 (1995) Soil quality - Sampling 

ISO 10381-6 (1993) Soil quality - Sampling - Part 6: Guidance on the collection, handling 
and storage of soil for the assessment of aerobic microbial processes in the laboratory 



1 Soil Sampling and Storage 45 

ISO 11266 (1994) Soil quality- Guidance on laboratory testing for biodegradation of organic 

chemicals in soil under aerobic conditions 
ISO 11277 (1998) Soil quality - Determination of particle size distribution in mineral soil 

material - Method by sieving and sedimentation 
ISO 11464 (1994) Soil quality - Pretreatment of samples for physico-chemical analysis 
ISO 1 5473 (2002) Soil quality - Guidance on laboratory testing for biodegradation of organic 

chemicals in soil under anaerobic conditions 
ISO 15799 (2003) Soil quality - Guidance on the ecotoxicological characterization of soils 

and soil materials 
Lund V, Goks0r J (1980) Effects of water fluctuations on microbial mass and activity in soil. 

Microbial Ecol 6:115-123 
Weinfurtner K, Koerdel W, Schlueter C (2002) Probenahmerichtlinie fur eine Kryo- 

lagerung von Bodenproben. Jahrestagungen GDCh-Fachgruppe Umweltchemie und 

Okotoxikologie / SETAC-GLB, Braunschweig 2002 



2 



Determination of Chemical 
and Physical Soil Properties 

Berndt-Michael Wilke 



2.1 

Soil Dry Mass and Water Content 

■ Introduction 

Objectives. Measures of soil water content and dry mass are needed in prac- 
tically all types of soil studies, e.g., determination of water holding capacity, 
plant available water, infiltration, pore size distribution, permeability. With 
respect to soil microbial processes and biological soil remediation, determi- 
nation of optimum water content for measurement of microbial parameters 
and activity, as well as determination of soil permeability for estimation of 
the success of in situ remediation, is of essential importance. 

Principle. Soil samples are dried at 105 ± 5°C until mass constancy is 
reached. The differences in masses before and after drying are a measure 
for the water content of soils. The water content is calculated on gravimetric 
(gwater/^soii) o r o n volumetric basis (cm 3 water /cm 3 SO ii). The method described 
below can be used for disturbed and undisturbed (sampling of soil using 
coring sieves) soil samples. It is a direct laboratory measurement. The 
procedure described can be used for the determination of dry mass on 
a mass basis (ISO 11465 1993). 

Theory. Under natural conditions all soils contain water. The amount of 
water can be very low in air-dried soils. As a convention the total water 
content and dry mass of soils are measured after drying at 105 °C (ISO 
1 1465 1993). Thus, the water content of a soil is given as percent by weight 
or volume of oven-dried soil. Water which is removed at higher tempera- 
tures is not included in the definition of soil water. The soil water content 
can be determined with direct and indirect methods. Direct methods are 
more precise but time consuming. Indirect methods are mainly used for 
continuous determination of water contents in the field. The most appro- 
priate indirect method is the time domain reflectometry (TRD) method 
(Topp et al. 2000). The optimum water content for microbial processes is in 



Berndt-Michael Wilke: Institute of Ecology, Berlin University of Technology, Franklinstrafte 
29, 10587 Berlin, Germany, E-mail: bmwilke@tu-berlin.de 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



48 B.-M. Wilke 

the range of 40-60% of maximum water-holding capacity (WHC, Sect. 2.2), 
or corresponds to the water content that is held in soil at suction pressures 
of -0.01 to -0.031 MPa. 

■ Equipment 

• Drying oven, thermostatically controlled with forced air ventilation and 
capable of maintaining a temperature of 105 ± 5 °C 

• Desiccator with an active drying agent 

• Analytical balance, accuracy 1 mg 

• Container (moisture box, 25-100 mL) with lid, made of waterproof ma- 
terial that does not adsorb moisture, capacity 25-100 mL for air-dried 
soil samples and at least 100 mL for field-moist soil samples 

• Spoon 

■ Procedure 

Air-Dried Soil Samples 

1. Dry container with lid at 105 ± 5 °C and then cool it, with the lid closed, 
in a desiccator for at least 45 min. Determine the mass (m ) of the closed 
container with an accuracy of ± 1 mg. 

2. Transfer 10-15 g of air-dried soil to this container using a spoon. 

3. Determine the mass (mi) of the closed container and soil with an accu- 
racy of ±1 mg. 

4. Dry the container and soil in an oven at 105 °C until constant mass is 
achieved. Dry the lid at the same time. 

5. Cool the container with the lid closed in a desiccator for at least 45 min. 

6. Remove the container from the desiccator and immediately determine 
the mass (m 2 ) of the closed container containing the oven-dried soil with 
an accuracy of ± 1 mg. 

Field-Moist Soil Samples 

1. Place the soil on a clean surface that does not absorb moisture (e.g., 
a glass plate) and mix well. Remove particles with a diameter > 2 mm. 

2. Dry container with lid at 105 ± 5 °C and then cool it, with the lid closed, 
in desiccator for at least 45 min. Determine the mass (m ) of the closed 
container with an accuracy of ± 1 mg. 

3. Transfer 30-40 g of soil to this container using a spoon. 



2 Determination of Chemical and Physical Soil Properties 49 

4. Determine the mass (mi) of the closed container and soil with an accu- 
racy of ±10 mg. 

5. Dry the container and soil in an oven at 105 °C until constant mass is 
achieved. Dry the lid at the same time. 

6. Cool the container with the lid closed in a desiccator for at least 45 min. 

7. Remove the container from the desiccator and immediately determine 
the mass (m 2 ) of the closed container containing the oven-dried soil with 
an accuracy of ± 10 mg. 

■ Calculation 

Calculate the dry mass content ( w dm ) or water content ( w H2 o ) on a dry mass 
basis expressed as percentages by mass to an accuracy of 0.1% (m/m) using 
the following equations: 

m 2 - m / 

w dm = — x 100 (2.1) 

mi - m 

mi - m 2 , 

w H0 = — x 100 (2.2) 

m 2 -m 

m mass of the empty container with lid (g) 

mi mass of the container with air-dried soil or field-moist soil (g) 

ra 2 mass of the container plus oven-dried soil (g) 

■ Notes and Points to Watch 

• With contaminated soil samples, special measures must be taken. Avoid 
any contact with the skin. Special measures must be taken during the 
drying process in order to prevent contamination of the laboratory at- 
mosphere. The procedures must be performed as quickly as possible to 
prevent evaporation. 

• In general decomposition of organic material can be neglected at tem- 
peratures up to 105 °C. However, for soil samples with a high organic 
matter content (> 10% m/m) the method of drying should be adapted 
by drying to a constant mass at 50 °C. 

• Some minerals similar to gypsum lose chemically combined water at 
a temperature of 105 °C. 

• If volatile organic substances are present, the method will not give a re- 
liable determination of the water content. 



50 B.-M. Wilke 

2.2 

Water-Holding Capacity 

■ Introduction 

Objectives. Microbiological laboratory tests (e.g., respiration measure- 
ments ISO 16072 2002; nitrogen mineralization ISO 14238 1997; biodegra- 
dation ISO 11266 1994) are carried out under optimum water conditions. 
These range from 40-60% of soil WHC. In order to adjust the optimum 
water content of a given soil the maximum WHC has to be determined. 

Principle. A cylinder with a perforated base is filled with soil, capped and 
immersed in water and drained. The quantity of water taken up by the 
soil is determined by weighing, drying to constant mass at 105 °C, and 
reweighing. 

Theory. Microbial transformations in soils are moisture dependent. Mois- 
ture must be adequate for decomposition to proceed. High moisture levels 
reduce activities of aerobic microorganisms due to a deficiency of oxygen. 
Therefore, the soil moisture content is adjusted to optimum conditions 
in microbial laboratory experiments. For most aerobic processes it ranges 
from 40 to 60% of WHC. Alternatively, the water content can be also ad- 
justed by means of pore water pressure. Water contents of 40-60% of WHC 
equal 0.01-0.03 MPa. 

■ Equipment 

• Cylinder (glass, plastic, metal; with cap) of known volume, of about 
50-150 mm length and 50-100 mm in diameter with a perforated base 

• Water bath at room temperature 

• Tray with a drainage hole, containing wet, fine, quartz sand (20-50 mm) 

• Oven, capable of maintaining a temperature of 105 ± 2 °C 

• Beaker, 250 mL 

• Desiccator 

• Balance, accurate to ±0.01 g. 

■ Sample Preparation 

As rule fresh soil samples screened through a 2 mm sieve are used. 



2 Determination of Chemical and Physical Soil Properties 51 

■ Procedure 

Determination of Maximum WHC 

1 . Cover the perforated base of the cylinder with a filter paper and fill it with 
field-moist soil (three parallels per sample). Fill the soil in small portions 
and provide homogeneous spreading by gentle tapping of the cylinder. 

2. Submerge the cylinder in the water bath at room temperature with the 
water level lower than the soil surface. When the soil is moistened to the 
surface, lower the cylinder to the soil surface and leave it in this position 
overnight. 

3. Remove the cylinder from the water and place the capped cylinder on 
the tray of sand and allow to drain. Capping of the cylinder is crucial to 
avoid evaporation of water. 

4. Weigh the cylinder hourly beginning after 3 h until constant weight is 
achieved. Remove the soil from the cylinder into a 250 mL beaker and 
dry it at 105 °C in an oven for 24 h (minimum). Cool the samples in 
a desiccator and weigh again. 

Adjustment of a Defined Water Content 

If the actual water content is lower than wanted, the soil is spread as a thin 
layer and the needed amount of water is evenly sprayed in small portions on 
the surface. The soil should be mixed thoroughly after each water addition. 
Reduction of soil volume (formation of aggregates) should be avoided 
during addition of water. 

In case of a higher actual water content the soil is dried at room temper- 
ature until the wanted moisture is reached. Drying of the soil surface has 
to be avoided by periodic mixing. 

The water content can also be adjusted by using a device (e.g., porous 
funnel apparatus) whereby the saturated soil can be drained stepwise to 
a known soil water (matric) potential. First, the field-moist soil is saturated 
on a ceramic plate. Subsequently the surplus of water is drained until the 
wanted water content is reached using a vacuum pump. 

■ Calculation 

Calculate the WHC using the following equation: 

WHC max (% dry mass) = — x 100 (2.3) 

m t - rab 

m s mass of beaker containing water saturated soil (g) 
m t mass of beaker containing oven-dried soil (g) 
mb mass of beaker (g) 



52 B.-M. Wilke 

2.3 

Bulk Density - Total Porosity 

■ Introduction 

Objectives. Determination of bulk density is a widely used soil parameter. 
Bulk density is needed for converting water percentage by weight to content 
by volume, calculating the porosity and void ratio when the particle density 
is known (Blake and Hartge 1986). It can by used to estimate the weight 
of a volume of soil too large to weigh and to calculate the total mass of 
a pollutant in a given soil volume. 

The bulk density gives a rough estimation of the aeration and permeabil- 
ity of a soil. The lower the bulk density, the higher is the permeability. Bulk 
density varies with structural conditions of the soil. Therefore, it is related 
to packing and often used as a measure for soil structure. In swelling soils 
(e.g., clay soils) it varies with soil water content (Hartge 1968). In these 
soils the bulk density obtained should be compared with the soil water 
content at the sampling time. There are three methods available for the 
determination of soil bulk density: core method, excavation method, and 
clod method. All methods are standardized (ISO DIS 11272 1998). 

Theory. Soil is a porous three-phase system composed of air, water, and 
solids. The relative distribution of these three components is important 
to understand the hydraulic properties of the soil. The dry bulk density 
is the ratio of oven-dried solids to volume of soil. It is expressed in SI 
units, e.g., g/cm 3 , kg/m 3 , or Mg/m 3 . It reflects the structural condition of 
the soil at given depth. Bulk densities of mineral soils may range from 
< 0. 8 to > 1.75 g/cm 3 (Schlichting et al. 1995). The total porosity (S t ) can be 
calculated if the particle density (p p ) and the bulk density (f>b) are known, 
according to the following equation: 

St = 1 - (pi/ft) (2.4) 

(Danielson and Sutherland 1986). As a rule of thumb the density of quartz 
(p p = 2.65 g/cm 3 ) is used as particle density (p p ) of mineral soils. 

2.3.1 

Core Method 

Principle. This method is only applicable to stoneless and slightly stony 
soils. A cylindrical metal sampler is pressed or driven into the soil to the 
desired depth. It is carefully removed to preserve a known volume of sample 
as it existed in situ. The sample is dried in an oven at 105 °C and weighed. 



2 Determination of Chemical and Physical Soil Properties 53 

■ Equipment 

• Core sampler holder, thin walled metal cylinders with a volume of 
100-400 cm 3 , a steel cap for driving into the soil, and a driver 

• Oven, heated and ventilated, capable of maintaining a temperature of 
105 °C 

• Desiccator 

• Laboratory balance, capable of weighing to an accuracy of 1/1,000 of the 
measured value 



■ Procedure 

1. Drive or press the core sampler into either a vertical or a horizontal soil 
surface enough to fill the sampler but not so firmly as to compress the 
soil in the confined space of the sampler. 

2. Carefully remove the sampler and its contents to preserve the natural 
structure, and trim the soil extending beyond each end of the sample 
holder with a straight-edged knife or a sharp spatula. The soil sample 
volume thus established is the same as the volume of the sample holder. 

3. Take at least six core samples from each soil horizon. 

4. Place the holders containing the sample in an oven at 105 °C until con- 
stant mass is achieved (minimum 48 h). 

5. Remove the samples from the oven and allow them to cool in the desic- 
cator. 

6. Weigh the samples immediately after removal from the desiccator (m t ). 
Control mass is reached when the differences in successive weighing of 
the cooled sample, at intervals of 4 h, do not exceed 0.01% of the original 
mass of the sample. 

■ Calculation 

The dry bulk density is calculated using the following equations: 

Qh = — (2.5) 

m& = m t — m s (2.6) 

£>b bulk density (g/cm 3 ) 

ma mass of the core sample dried at 105 °C minus mass of the core sample 
holder (g) 



54 B.-M. Wilke 

V volume of the core sample holder (cm 3 ) 

m t mass of the sample holder plus soil sample dried at 105 °C (g) 

m s mass of the empty core sample holder (g) 

■ Notes and Points to Watch 

• Swell/shrink soils (e.g., clays, muds, peats) change their bulk density 
with changing water content. Such soils should be sampled in a moist 
state (e.g., field capacity). In addition they should be sampled in a wet 
state (water saturation) and a dry state. 

• If bulk density and water content (Sect. 2.1) are the only parameters of 
interest, it is not necessary to keep the samples in their holders. A single 
core sample holder can be reused if each sample is transferred to another 
container. 

• The undisturbed samples in the core sample holder can be also used for 
other measurements such as pore-size distribution (Sect. 2.5), conduc- 
tivity, or water retention. 

• It is normally worthwhile to combine a measurement of the water content 
with a measurement of bulk density. In this case, it is necessary to 
transport the samples without allowing loss of water by evaporation and 
to begin the laboratory operations by weighing the fresh sample. 



2.3.2 

Excavation Method 

Principle. Dry bulk density is determined by excavating a quantity of soil, 
drying and weighing it, and determining the volume of the excavation by 
filling it with sand. The method is applicable to soils containing gravel 
and/or stones. 

■ Equipment 

• Earth-digging equipment, e.g., spade with sharp-edged blade 

• Sampling equipment (flat blade spade, knife, pick, spade chisel, hammer) 

• Equipment for collecting and cleaning (plastic sheet, brush, heat-resistant 
plastic bags or containers) 

• Plastic film, thin, flexible but stable 



2 Determination of Chemical and Physical Soil Properties 55 

• Equipment for spreading sand, including funnel with a gauging rod 
(falling height beneath the funnel mouth should be 5 cm), graduated 
cylinder of 1 dm 3 capacity 

• Dry, graded sand of known volume, particle diameter 500-700 yam 

• Balance capable of weighing 1 g 

• Oven, heated and ventilated, capable of maintaining a temperature of 
105 °C 

• Vacuum desiccator with self indicating desiccant 

• Sieve with 2-mm mesh size 

■ Procedure 

Field 

1. Level off the soil surface with the straight metal blade (Fig. 2.1a). 

2. Dig a hole in the leveled soil having a representative content of larger 
gravel and stones (volume 20 dm 3 containing 30% stones) avoiding com- 
paction of sides (Fig. 2.1b). 

3. Put the excavated soil in bags or containers for laboratory analysis. (Large 
nonporous stones such as granite can be separated in the field, cleaned 
with a stiff brush, and weighed on a field balance). 

4. Line the hole with a plastic film. 

5. Fill the hole to excess with a known volume of sand from a height of 5 cm 
using the funnel (Fig. 2.1c). Level the surface with the blade without 
packing down. 

6. Pour the excess sand into the graduated measuring cylinder and read 
the volume (Fig. 2. Id). The difference from the initial volume of sand is 
the volume V in the hole. 

Laboratory 

1. Determine the mass of the moist excavated soil (in g) with a balance 

2. Separate the stones and gravel from the fine soil with a 2-mm sieve and 
weigh them on a balance (m xw ). 

3. Dry the stones and the gravel in the oven at 105 °C and weigh them after 
cooling on the laboratory balance (m x ). 

4. Determine the water content of the fine soil (< 2 mm) by drying a rep- 
resentative sample (5-10 g) of known mass in the oven (105 °C) until 



56 



B.-M. Wilke 



^ 



^ 







a 



^ 





Fig. 2.1. Excavation method, field procedure (adapted from ISO 11272 1998). a Level off soil 
surface; b dig a hole; c fill with sand; d remove excess sand and measure its volume 

constant mass is reached. Remove the sample from the oven and cool it 
in the desiccator. Weigh the sample on a laboratory balance. Calculate 
the water content (w) as a mass ratio of the moist sample. 



Qh = 



m x - m tp 



■ Calculation 

The bulk density of the soil layer is calculated using the following equations: 

(2.7) 

(2.8) 
m w = m pw x m tw (2.9) 

(2.10) 



V 



™tp ~ wip w ■ ■ trixw ' ' wi w 



TH-Ua; — ffltiAA; ' ' ffl 



■tw 



pw 



l xw 



£>b bulk density (g/cm 3 ) 

m x mass of stones and dry gravel (g) 

m tp mass of the dry fine soil (g) 



2 Determination of Chemical and Physical Soil Properties 57 

V volume of the hole (cm 3 ) 

m pw mass of the excavated moist soil (g) 

m w mass of the water from excavated fine soil (g) 

w water content of the excavated moist fine soil (g/g oven-dried soil) 

m tw mass of the moist fine soil (g) 

m xw mass of moist gravel and stones (g) 

■ Notes and Points to Watch 

• Holes should have smooth, rounded walls. 

• Protruding stones should be included in the sample 

• A heavy pair of scissors can be used to cut roots at the wall surface. 

2.3.3 

Clod Method 

Principle. The bulk density of clods, or coarse peds, is calculated from 
their mass and volume. The volume is determined by coating the clod 
with a water-repellent substance and by weighing it first in air, then again 
while immersed in a liquid of known density, making use of Archimedes' 
principle. 

■ Equipment 

• Earth digging equipment (flat shovel, spade, pick) 

• Sampling equipment (small flat-bladed spade, knife, chisel, hammer) 

• Container of molybdenum sulfide (MoS 2 ) in heavy oil 

• Laboratory balance, capable of weighing suspended samples (Fig. 2.2) 

• Thermometer 



■ Procedure 

1. Separate clods or peds of about 50-200 cm 3 , trim off protrusions and 
cut off roots with scissors. 

2. Weigh soil clods or peds with a laboratory balance and coat them in oil. 



58 



B.-M. Wilke 



1 
2 



3 
4 




Fig. 2.2. Laboratory balance to determine the volume of clods by weighing in air and water. 
1 Compensating weights; 2 thin wire; 3 small container; 4 large container filled with water; 
5 balance 



3. Weigh the coated clod again once in air and once immersed in water. 

4. Measure the temperature of the water and determine its density 

5. To obtain a correction for the water content of the soil, break the clod, 
remove an aliquot of soil, and weigh it before and after drying in an oven 
atl05±2°C. 



■ Calculation 

1. The oven dry mass of the soil clods is calculated using the equation 



m& = ra/(l + w) 



(2.11) 



ma net mass of the oven-dried clod (g) 
m net mass of the moist clod in air (g) 
w water content of the subsample (g of water/g of oven-dried soil) 



2. Calculate the bulk density of the dried clod using the equation 



mass 



Pb = 



pw x m d 



volume m- m w + m (p - £>w) 



(2.12) 



05 bulk density of oven-dried sample (g/cm 3 ) 

(? density of the coating oil (g/cm 3 ) 

p w density of water at temperature of determination (g/cm 3 ) 

ma oven-dried mass of soil sample, i.e., clod or ped (g) 



2 Determination of Chemical and Physical Soil Properties 59 

m mass of soil sample in air (g) 

m w mass of soil sample plus coating in water (g) 

m mass of coating in air (g) 

■ Notes and Points to Watch 

• The clod method gives usually higher bulk density values than the other 
methods as it does not take interclod space into account. 

• Clods on or near the soil surface are likely to be unrepresentative as these 
are often formed by packing or plowing. 

• Several other substances have been used to seal the clods against water 
including Saran solution (Dow Chemical, Rolling Meadows, IL, USA), 
paraffin, and wax mixtures. 

• See also Blake and Hartge (1986). 

2.4 

Water Retention Characteristics - Pore Size Distribution 

■ Introduction 

Objectives. The spaces between soil particles are known as the soil pores. 
They are filled either with soil-air or water (soil solution) depending on the 
pore size and the water saturation of the soil. With respect to their equiv- 
alent diameter, soil pores can be divided into wide coarse (> 50 yam), tight 
coarse (10-50 yam), medium (0.2-10 yam) and fine (< 0.2 yam). Pore sizes 
were assigned in accordance with adaptation to the water content at char- 
acteristic matric pressures. Equivalent diameters of 50 and 10 yam comply 
with the water content of soils at field capacity (6 and 30 kPa), 0.2 yim with 
the water content at the permanent wilting point (1,500 kPa). The range of 
water available to plants and microorganisms is between field capacity and 
the permanent wilting point. Water stored at matric pressures > 1,500 kPa 
is neither accessible to fine plant roots nor to microorganisms. The pore size 
distribution of a given soil depends on its density and texture. Thus, it in- 
fluences its aeration, permeability, transport of chemicals dissolved in soil 
water, and the water- retention characteristics of a soil. Direct evaluation of 
the size, configuration, and distribution of soil pores is impossible due to 
their extremely complicated nature. However, the size distribution can be 
measured by determination of water content at different matric pressures. 
Besides providing an assessment of the equivalent pore size distribution 
(e.g., identification of coarse, medium, and fine pores), the results using 
the methods to be described can be used for other purposes, for example: 



60 B.-M. Wilke 

• For assessment of the water retention characteristics 

• To determine water content at specific matric pressures (e.g., for micro- 
bial degradation studies) 

• To ascertain the relationship between the negative matric pressures and 
other soil physical properties (e.g., hydraulic conductivity, thermal con- 
ductivity) 

• To determine the drainable pore space (e.g., pollution risk assessment) 

• To determine indices for plant- available water in the soil (e.g., for irri- 
gation purposes) 

Principle. Undisturbed soil samples (soil cores) are used for the measure- 
ment at the high matric pressure range 0-100 kPa. The samples are satu- 
rated with de-aerated water or calcium sulfate solution (0.005 mol/L) and 
subsequently drained using sand, kaolin, or ceramic suction tables (for 
pressures from to 20 kPa) and pressure plate extractors (for determina- 
tion of pressures from -5 to -1,500 kPa). At equilibrium status, soil samples 
are weighed, oven dried and reweighed to determine the water content. The 
results are given either as volume fraction or mass ratio. The differences in 
volume fractions at different suction pressures give the pore volume (e.g., 
medium pores in vol%), the differences in mass fractions give the water 
content retained in these pores. Two standardized (ISO 11274 1998) meth- 
ods are described, namely use of sand, kaolin, or ceramic suction tables 
for determination of water contents at pressures of to -50 kPa, and use of 
pressure plates for determination of pressures from -5 to -1,500 kPa. 

Theory. Soil water content and matric pressure are related to each other. At 
zero matric pressure the soil is saturated and all pores are filled with water. 
As the soil dries matric pressure decreases and pores will empty according 
to their equivalent diameter. Large coarse pores (> 50 p.m) will drain at 
a matric pressure of > -6kPa, tight coarse pores (10-50 }im) at -6 to 
30 kPa, medium pores at -30 to -1,500 kPa, and fine pores at < -1,500 kPa. 

■ Sampling 

1. It is essential that undisturbed soil samples be used for measurement 
at the matric pressure range to -lOOkPa, since soil structure has 
a strong influence on water- retention properties. Use either undisturbed 
cores or, if appropriate, individual peds for low matric pressure methods 
(< -100 kPa). Soil cores shall be taken in a metal or plastic cylinder of 
a height and diameter such that they are representative of the natural 
soil variability and structure. The dimensions of samples taken in the 
field are dependent on the texture and structure of the soil and the test 



2 Determination of Chemical and Physical Soil Properties 61 

Table 2.1. Recommended sample sizes (height x diameter) for the different test methods 



Test method 




Structure 






Coarse 


Medium 


Fine 


Suction table 
Pressure plate 


50 x 100 mm 


40 x 76 mm 
10 x 76 mm 


24 x 50 mm 
10 x 50 mm 



method which is to be used. Table 2.1 gives guidance on suitable sample 
sizes for the different methods and soil structure. 

2. To ensure minimal compaction and disturbance to structure, take soil 
cores carefully, either by hand pressure in suitable material or by using 
a suitable soil corer. Take a minimum of three representative replicates for 
each freshly exposed soil horizon or layer; more replicates are required 
in stony soils. Dig out the cylinder carefully with a trowel, roughly trim 
the two faces of the cylinder with a knife. If necessary adjust the sample 
within the cylinder before fitting lids to each end, and label the top clearly 
with the sample grid reference, the direction of the sampling (horizontal 
or vertical), the horizon number, and the sample depth. 

3. Wrap the samples (e.g., in plastic bags) to prevent drying. Wrap ag- 
gregates (e.g., in aluminum foil or plastic film) to retain structure and 
prevent drying. Alternatively, excavate undisturbed soil blocks measur- 
ing approx. 30 cm 3 in the field, wrap in metal foil, wax (to retain structure 
and prevent drying), and take to the laboratory for subdivision. Store the 
samples at 1-2 °C to reduce water loss and suppress biological activity 
until they can be analyzed. Treat samples having obvious macrofaunal 
activity with a suitable biocide, e.g., 0.05% copper sulfate solution. 



■ Sample Preparation 

1. To prepare samples for water- retention measurements at pressures great- 
er than -50kPa, trim undisturbed cores flush with the ends of the con- 
tainer and replace one lid with a circle of polyamide (nylon) mesh (or 
similar close-weave material or paper if the water- retention character- 
istic is known) secured with an elastic band. The mesh will retain the 
soil sample in the cylinder and enable direct contact with the soil and 
the porous contact medium. Avoid smearing the surface of clayey soils. 
Remove any small projecting stones to ensure maximum contact and 
correct the soil volume if necessary. Replace the other lid to prevent 
drying of the sample by evaporation. Prepare soil aggregates for high 
matric pressure measurements by leveling one face and wrapping other 



62 B.-M. Wilke 

faces in aluminum foil to minimize water loss. Disturbed soils should be 
packed into a cylinder with a mesh attached. Firm the soil by tapping 
and gentle pressure to obtain a specified bulk density. 

2. Weigh the prepared samples. Ensure that the samples are brought to 
a pressure of less than the first equilibration point by wetting them, if 
necessary, by capillary rise, mesh side or leveled face down on a sheet of 
foam rubber saturated with de-aerated tap water or 0.005 mol/L calcium 
sulfate solution. Weigh the wet sample when a thin film of water is seen 
on the surface. The time required for wetting varies with initial soil 
water content and texture. Soils are ideally field moist when the wetting 
is commenced. General guidelines for wetting times are: 

sand: 1-5 days 

loam: 5-10 days 

clay: 5-14 days or longer 

peat: 5-20 days. 

Very coarse pores are not water filled when the soil sample is saturated 
by capillary rise. 

2.4.1 

Determination of Soil Water Characteristics 

Using Sand, Kaolin, and Ceramic Suction Tables 

Principle. Suction tables are suitable for measurement of water contents at 
matric pressure from to -50 kPa. A negative matric pressure is applied to 
coarse silt or very fine sand held in a rigid watertight non-rusting container 
(a ceramic sink is particularly suitable). Soil samples placed in contact 
with the surface of the table lose pore water until their matric pressure is 
equivalent to that of the suction table. Equilibrium status is determined by 
weighing samples on a regular basis, and soil water content by weighing, 
oven drying, and reweighing. 

■ Equipment 

• Large ceramic sink or other watertight, rigid, non-rusting container with 
outlet in base (dimensions about (50 x 70 x 25 cm) and close-fitting cover 

• Tubing and connecting pieces to construct a draining system for the 
suction table 

• Sand, silt, or kaolin, as packing material for the suction table (Com- 
mercially available graded and washed industrial sands with a narrow 



2 Determination of Chemical and Physical Soil Properties 



63 



particle size distribution are most suitable. The particle size distribu- 
tions of some suitable sand grades and the approximate suctions they 
can attain are given in Table 2.2. It is permissible to use other packing 
materials, such as fine glass beards or aluminum oxide powder, if they 
can achieve the required air entry values. Alternatively to sand, silt, or 
kaolin suction tables, ceramic plates can be used. 

Leveling bottle, stopcock, and 5-L aspirator bottle 

Tensiometer system (optional) 

Drying oven, capable of maintaining a temperature of 105 ± 2 °C 

Balance capable of weighing with an accuracy of 0.1% of the measured 
value 



■ Procedure 

1 . Prepare suction tables using packing material that can attain the required 
air entry values (Table 2.2). 

2. Prepare soil cores as described (see above). 

3. Weigh the cores and then place them on a suction table at the desired 
matric pressure. 

4. Leave the cores for 7 days. The sample is than weighed, and thereafter 
weighed as frequently as needed to verify that the daily change in mass of 
the core is less than 0.02%. The sample is than regarded as equilibrated. 



Table 2.2. Examples of sands and silica flour suitable for suction tables 



Type 



Coarse sand 



Medium 



Fine sand 



Silica flour 



Use 



Base of 

suction 

tables 



Surface of Surface of Surface of 

suction tables suction tables suction tables 

(5 kPa matric (11 kPa matric (21 kPa matric 

pressure) pressure) pressure) 



Typical particle 
size distribution 



Percent content 



> 600 um 


1 


1 


1 





200-600 um 


61 


8 


1 





100-200 jim 


36 


68 


11 


1 


63-100 um 


1 


20 


30 


9 


20-63 um 


1 


3 


52 


43 


<20um 








5 


47 



64 B.-M. Wilke 

5. Move the equilibrated sample to a suction table of a lower pressure or 
dry it in an oven at 105 ± 5 C. 

6. Samples which have not attained equilibrium should be replaced firmly 
onto the suction table and the table cover replaced to minimize evapo- 
ration from the table. 

■ Calculation 

Soils Containing <20% Stones (>2mm) 

1. Calculate the water content mass ratio at a matric pressure p m using the 
formula: 

w(p m ) = -^ (2.13) 

m d 

w(p m ) water content mass ratio at a matric pressure p m (g) 
m(p m ) mass of the soil sample at a matric pressure p m (g) 
ma mass of the oven-dried soil sample (g) 

2. Calculate the water content on a volume basis at matric pressure p m using 
the formula: 

/ x m (pm) - md 

e p m = -^r 1 — - (2.14) 

v y V x p w 

0(pm) water content mass ratio at a matric pressure p m 
(cm 3 water/ cm 3 soil) 

m(p m ) mass of the soil sample at a matric pressure p m (g) 

ma mass of the oven-dried soil sample (g) 

V volume of the soil sample (cm 3 ) 

p w density of water (g/cm 3 ) 

Conversion of Results to a Fine Earth Basis 

The stone content of a laboratory sample may not accurately represent the 
field situation. Therefore, conversion of data to a fine earth basis may be 
required. Conversion of results derived from suction methods to a fine 
earth basis (f) is required for soils containing stones (> 2 mm) according 
to the following equation: 

"• ■ Ay <2 - i5) 



2 Determination of Chemical and Physical Soil Properties 65 

0f water content of the fine earth expressed as a volume fraction 
6 S volume of stones, expressed as a fraction of total core volume 
6 t water content of the total soil, expressed as a volume fraction 

2.4.2 

Determination of Soil Water Characteristics 

by Pressure Plate Extractor 

Principle. Pressure plate extractors are suitable for measurement of water 
contents at matric pressure -5 to -1,500 kPa. Several small soil cores are 
placed in contact with a porous ceramic plate contained within a pressure 
chamber. A gas pressure is applied to the air space above the samples and 
soil water moves through the plate to be collected in a burette/measuring 
cylinder or similar collecting device. At equilibrium status, soil samples 
are weighed, oven-dried, and reweighed to determine the water content at 
the predetermined pressures. 

■ Equipment 

• Pressure chamber with porous ceramic plate 

• Sample retaining rings/soil cores with plastic discs or lids 

• Graduated burette 

• Air compressor (1.700 kPa), nitrogen cylinder, or other suitable pressur- 
ized gas 

• Pressure regulator and test gauge 

• Drying oven capable of maintaining a temperature of 105 ± 2.0 °C 

• Balance capable of weighing to ±0.01 g 

■ Procedure 

1. Take small soil cores of approx. 5 cm diameter and 5-10 mm in height 
in situ or from larger undisturbed cores. 

2. Place at least three replicates on a pre-saturated plate of appropriate 
bubbling pressure. 

3. Wet the samples by immersing the plate and the samples to a level just 
above the base of the core and waiting until a thin film of water can be 
seen on the surface of the sample. 



66 B.-M. Wilke 

4. Cover the bottom of the extractor with water to create a saturated 
atmosphere. 

5. Place a plastic disc lightly on top of each sample to prevent evaporation. 

6. To apply the desired pressure, remove excess water from the porous 
plate and connect the outflow tube to the burette via the connector in 
the chamber wall. The pressure is supplied via regulators and gauges 
from a nitrogen cylinder or by a mechanical air compressor. 

7. The pressure (from whatever source) should slightly exceed the lowest 
matric pressure required. 

8. Apply the desired gas pressure p, check for any gas leaks, and allow the 
samples to come to equilibrium by recording on a daily basis the volume 
increase in the burette. When this remains static, the samples have come 
to equilibrium; the matric pressure p m of the samples equals -p. 

9. To remove the samples, clamp the outflow tube to prevent a backflow 
of water, and release the air pressure. 

10. Weigh the samples plus sleeve immediately. 

11. Carry out sequential equilibration of the core at different pressures 
by removing and weighing the core at equilibrium, reinserting it, and 
resetting the pressure. 

12. Moisten the ceramic plate with a fine spray of water to re-establish 
hydraulic contact. 

13. When the last equilibrium has taken place, dry at 105 °C and determine 
the oven-dried mass of the soil plus sleeve. 



■ Calculation 

Stoneless Soils 

Calculate the water content volume fraction (0) using the formula: 

0{p m ) = ^l 1 ( 2 .1 6 ) 

Q(pm) water content mass ratio at a matric pressure p m (cm 3 water/cm 3 
soil) 

m(p m ) mass of the soil sample at a matric pressure p m (g) 

ma mass of the oven-dried soil sample (g) 



2 Determination of Chemical and Physical Soil Properties 67 

V volume of the soil sample (cm 3 ) 

p w density of water (g/cm 3 ) 

Stony Soils 

Samples containing any stones (> 2 mm) shall not form part of the pressure 
chamber or membrane sample since the sample volume is very small. After 
oven-drying, determine the volume of stones in the original soil core from 
a field measurement and make a correction to convert 0f values to total 
soil (0 t ). 

e t = e f (i-e s ) (2.17) 

0f water content of the fine earth in the pressure vessel at equilibrium 
expressed as volume fraction 

6 S volume of stones, expressed as a fraction of total core volume 

6 t water content of the total soil, expressed as a volume fraction 

For a soil containing a volume fraction of non porous stones of 0.05 the 
water content is: 

t = f x 0.95 (2.18) 

Evaluation of Results: Pore Size Distribution 

Pore volumes of coarse, medium, and tight pores in vol% of total soil 
volume can be calculated as follows: 

Large Coarse Pores (Equivalent Diameter > SOjjm) 

V\c V = (0pmO-0pm-6) * 100 (2.19) 

Vi C p volume of large coarse pores (% of total soil volume) 
0pmo volumetric water content at water saturation (p m = kPa) 
0pm-6 volumetric water content at a matric pressure of p m = -6 kPa 

Tight Coarse Pores (Equivalent Diameter 10-50 pm) 

V tcv = (0 pm -6 " 0pm-3o) x 100 (2.20) 

V t cp volume of large coarse pores (% of total soil volume) 
0pm-6 volumetric water content at water saturation (p m = -6 kPa) 
0pm-3o volumetric water content at a matric pressure of p m = -30 kPa 



68 B.-M. Wilke 

Medium Pores (Equivalent Diameter 0.2-10 pm) 

V mV = {0-30 - 0pm-15Oo) X 100 ( 2 - 21 ) 

V mp volume of large coarse pores (percent of total soil volume) 
0pm-3o volumetric water content at water saturation (p m = -30 kPa) 
0pm-i5oo volumetric water content at a matric pressure of p m = -1,500 kPa 

Fine Pores (Equivalent Diameter < 0.2 pm) 

Vfp = 0pm-15OO x 100 (2.22) 

Vfp volume of fine pores (% of total soil volume) 

0pm-i5oo volumetric water content at a matric pressure of p m = -1,500 kPa 

■ Notes and Points to Watch 

• If a containing sleeve is used, it should be weighed and the mass deducted 
from the total mass of the soil core to give m(p m ). 

• If stones are porous, carry out separate water retention measurements 
and correct fine earth values according to their volume. 



2.5 
Soil pH 

■ Introduction 

Objectives. Soil pH is one of the most indicative measurements of the soil 
chemical properties. All (bio)chemical reactions in soils are influenced by 
proton (H + ) activity, which is measured by soil pH. Values of pH of most 
natural soils (measured in 0.01 M CaCl 2 ) vary between < 3. 00 (extremely 
acid) and 8.00 (weakly alkaline). Solubility of various compounds in soils 
is influenced by soil pH (e.g., heavy metals) as well as by microbial activ- 
ity and microbial degradation of pollutants. The optimum pH values for 
pollutant-degrading microorganisms range from 6.5 to 7.5 (Kastner 2001). 
Determination of soil pH is standardized in ISO DIS 10390 (2002). 



2 Determination of Chemical and Physical Soil Properties 69 

Principle. A pH measurement is normally made by either a colorimetric or 
an electrometric method. The former involves suitable dyes or acid-base 
indicators. Indicator strips can be used for rough estimation of soil pH. 
Normally, pH values of soils are measured by means of a glass electrode 
in a soil solution slurry that contains a fivefold volume of water containing 
lMKClor0.01MCaCl 2 . 

Theory Soil pH is a measure of the activity of ionized H (H + , H 3 + ) and 
defined as the negative logarithm of the H + /H 3 + ion activity in mol/L. 
Soil acidity results from soluble acids in the soil solution, e.g., organic acids 
and carbonic acid. Further acidic cations in the soil solution are Al 3+ and 
Fe 3+ . Al 3+ ions exists in water as an [A1(H 2 0) 6 ] 3+ complex which dissociates 
intoH 3 + ions according to [A1(H 2 0) 6 ] 3+ + H 2 ^ [A1(H 2 0) 5 ] 2+ + H 3 + 
(pK a = 5. 0). A stronger cationic acid producer is Fe 3+ (pK a = 2. 2), which 
due to the low solubility of iron oxides only exists below pH 3. 

Soil pH is influenced by various factors, namely, the nature and type of 
inorganic and organic constituents (that contribute to soil acidity), the 
soil/solution ratio, the salt or electrolyte content, and the C0 2 partial 
pressure. A pH measurement in water includes easily dissociated pro- 
tons while 0.01 M CaCl 2 and 1 M KC1 solutions also mobilize exchangeable 
H + . They are used to simulate soil solutions of arable soils (CaCl 2 ) and 
forest soils (KC1) in temperate humid climates. Values of pH measured at 
constant salt concentrations reflect seasonal variations to a lower degree 
(Page et al. 1982); and those measured in 0.01 M CaCl 2 are 0. 6 ± 0. 2 units 
lower than pH H2 o values, because H + and Al 3+ ions are partly exchanged 
byCa 2+ . 

■ Equipment 

• Shaking or mixing machine 

• pH meter with slope adjustment and temperature control (in case of pH 
values > 10, an electrode specifically designed for that range is to be 
used) 

• Glass electrode and a reference electrode or a combined electrode of 
equivalent performance 

• Thermometer capable of measuring to the nearest 1 °C 

• Sample bottle (50 mL) made of borosilicate glass or polyethylene with 
a tightly fitting cap 

• Spoon of known capacity (at least 5.0 mL) 



70 B.-M. Wilke 

■ Reagents 

• Water with a specific conductivity not higher than 0.2 mS/m at 25 °C and 
apH> 5.6 

• Potassium chloride solution (KC1 1 mol/L) 

• Calcium chloride solution (CaCl 2 0.01 mol/L) 

• Solution for the calibration of the pH meter 

• Buffer solution, pH 4.00 at 20 °C: dissolve 10.21 g of potassium hydrogen 
phthalate (C6H5O4K, dried at 1 10-120 °C for 2h before use) in water 
and dilute to 1,000 mL at 20 °C. 

• Buffer solution, pH 6.88 at 20 °C: dissolve 3.39 g of KH 2 P0 4 and 3.53 g of 
Na 2 HP0 4 in water and dilute to 1,000 mL at 20 °C. 

• Buffer solution, pH 9.22 at 20 °C: dissolve 3.80 g of Na 2 B 4 7 x 10H 2 Oin 
water and dilute to 1,000 mL at 20 °C. The buffer solutions are stable for 
1 month when stored in polyethylene bottles. Alternatively, commercially 
available buffer solutions may be used. 

■ Sample Preparation 

Use the fraction of particles of air-dried soil or soil dried at temperatures 
< 40 °C and passed through a square-hole sieve with 2-mm mesh size. 
Alternatively, field-moist soil passed through a 2-mm sieve can be used. 

■ Procedure 

1. Take a representative test portion of at least 5 mL from the soil sample 
using the spoon. 

2. Place the test portion in the sample bottle and add five volumes of water, 
potassium chloride solution, or calcium chloride solution. 

3. Shake or mix the suspension for 60 ± 10 min using a mechanical shaker 
(never longer than 3h). The stirring should be at a rate that achieves 
a homogenous soil suspension. Entrainment of air should be avoided. 

4. Calibrate the pH meter as prescribed in the manufacturer's manual using 
the buffer solutions. 

5. Adjust the pH meter as indicated in the manufacturer's manual. Measure 
the temperature of the suspension and take care that the temperature of 
the buffer and the soil solution does differ more than 1 °C. Measure the 
pH in the suspension while or immediately after being stirred. Read the 
pH after stabilization is reached. Record the pH values to two decimals. 



2 Determination of Chemical and Physical Soil Properties 71 

■ Notes and Points to Watch 

• Drying may influence the pH of soils, especially those containing sulfides. 
In such soils drying will lower the pH substantially. 

• In calcareous soil samples the pH depends on calcium ion activity and 
C0 2 partial pressure (pC0 2 ), and also on the quality of the laboratory 
air (Schlichting et al. 1995). 

• If a swinging-needle pH meter is used, the second decimal place should 
be estimated (ISO DIS 10390 2002). 

• In samples with a high content of organic material (e.g., peat soils, 
pot soils) the suspension effect can play a role. In calcareous soils it is 
possible for carbon dioxide to be adsorbed by the suspension. Under 
these circumstances it is difficult to reach equilibrium pH values (ISO 
DIS 10390 2002). 

• Magnetic stirring of the suspension is not suitable since this can affect 
the reading of pH. 

• pH indicator strips may be used for rough estimations. 

2.6 

Soil Organic Matter - Soil Organic Carbon 

■ Introduction 

Objectives. Soil organic matter (SOM) is one of the most important indi- 
cators of soil quality. It influences many soil properties including nutrient 
supply (mainly N, P, S), cation exchange capacity, adsorption of pollu- 
tants, infiltration and retention of water, soil structure, and soil color, most 
of which in turn affect soil temperature. SOM consists of microbial cells, 
plant and animal residues at various stages of decomposition, stable humus 
(humic acids, humins) synthesized from residues by microorganisms, and 
highly carbonized compounds (e.g., charcoal, graphite, coal; Nelson and 
Sommers 1996). The term humus is used synonymously with SOM; that is, 
it denotes all organic material in the soil. Organic material is essential as 
a nutrient source for all heterotrophic soil organisms, which in turn hold 
a key position in the processes of humification and mineralization of humic 
substrates that lead to the production of stable humus, degradable organic 
compounds, and carbon dioxide (Forster 1995a). There is often a direct 
relationship between the organic carbon contents of soils and microbial 
biomass and activity. Several methods are available for the determination 
of SOM in soils. Most often SOM content of soils is determined by carbon 



72 B.-M. Wilke 

analysis. Two methods are described in this Section, namely dry combus- 
tion and loss on ignition (LOI). 

Theory. Carbon is the chief element (48-58%) in SOM. Therefore, organic 
C determination is used as a basis of SOM estimates in soils. Based on 
the assumption that SOM contains 58% organic C, a conversion factor of 
1.724 has been proposed for the conversion of organic C content to SOM 
(humus content) of soils (Nelson and Sommers 1996). C content of soil 
can be determined by wet and dry combustion techniques. If inorganic C 
is also extracted, corrections have to be made for the inorganic portion. 
This can be done either by destruction of inorganic C prior to C analysis 
or by separate measurement and subtraction of inorganic C from total 
C content. Wet digestion procedures are based on oxidation of organic C 
compounds by Q^Oy". Because of the high toxicity of Cr(VI) compounds, 
this method should not be used. Dry combustion techniques are based on 
heating the soil gradually up to > 900 °C and subsequent measurement of 
evolved C0 2 trapped in a suitable reagent and determined titrimetrically 
or gravimetrically. There are also other measuring devices in use (see below 
and ISO 10694 1995). A simple technique for the estimation of SOM is the 
LOI method that was standardized in Germany under DIN 19684-3 (1977). 

2.6.1 

Dry Combustion Method 

Principle. The soil sample is gradually heated in a stream of purified oxy- 
gen to > 900 °C. Organic and inorganic soil carbon is converted to C0 2 . 
The C0 2 evolved is measured by titrimetry, gas chromatography, infrared 
spectrometry, or gravimetry. In the presence of carbonates, the samples 
are pretreated with HC1. If the carbonate content is known (determination 
according to ISO 10693 1995), the organic carbon can be calculated. Soils 
with pH(CaCl 2 ) < 6. 5 are unlikely to contain carbonates! 

■ Equipment 

• Analytical balance, accuracy 0.1 mg, or microbalance, accuracy 0.01 mg. 

• Apparatus for determination of total organic carbon by dry combustion 
at a temperature of > 900 °C equipped with an appropriate C0 2 de- 
tector. The following detection devices are currently available: titrime- 
try, gravimetry, gas chromatography, conductometry, and infrared spec- 
troscopy. Some of the devices are able to measure separately inorganic 
and organic carbon, others also measure total C and N contents (CN 
analyzer) 



2 Determination of Chemical and Physical Soil Properties 73 

• Crucibles made of porcelain, quartz, silver, tin, or nickel of different size; 
crucibles made of tin and nickel are not acid resistant. 



■ Reagents 

• Distilled or demineralized water with an electric conductivity of < 
0.2mS/mat25°C 

• Reagents for calibration, e.g., acetanilide (C 8 H 9 NO); atropine 
(Q7H23NO3); calcium carbonate (CaC0 3 ); graphite powder for spec- 
troscopy (C); sodium hydrogen phthalate (C 8 H 5 K0 4 ) 

• HC1 (4 mol/L) 

■ Sample Preparation 

Use air-dried, sieved (< 2 mm) soil. 

■ Procedure 

1. Weigh out mi g of the air-dried sample or subsample into a crucible. The 
amount for analysis depends on carbon content and on the apparatus 
used! 

2. Carry out the analysis according to the manufacturer's manual. 

3. Soils containing carbonates should be pretreated as follows: add an excess 
of HC1 to the crucible containing a weighed quantity of air-dried soil and 
mix. Wait 4 h and dry the crucible for 16 h at a temperature of 60-70 °C. 
Then carry out the analysis in accordance to the manufacturer's manual. 
The quantity of HC1 depends on the weight of the subsample and its 
carbonate content. In all cases an excess of acid should be added! 

■ Calculation 

Organic Carbon Content 

The total carbon content is calculated according to the following equation: 

m 2 100 + w H? o 

w ct = 1000 x — x 0.2727 x — (2.23) 

mi 100 

wet total carbon content on the basis of oven-dried soil (g/kg) 

mi mass of the test portion (g) 

m 2 mass of carbon dioxide released by the soil sample (g) 



74 B.-M. Wilke 

0.2727 conversion factor for C0 2 to C 

Wh 2 o water content expressed as a percentage by mass on a dry mass 
basis (Sect. 2.1) 

Organic Matter Content 

The organic matter content of the soil sample can be calculated using the 
following equation: 

Wom = / X Wcorg (2.24) 

w om organic matter content of the soil on the basis of oven-dried soil 
(g/kg) 

wcorg organic carbon content of the soil on the basis of oven-dried soil 
(g/kg) 

/ conversion factor 

2.6.2 

Loss On Ignition Method (LOI) 

Principle. The LOI method is based on ignition (550 ± 25 °C) of a dried 
(105 °C) soil sample until mass constancy is achieved. The SOM content is 
calculated from the mass difference before and after heating. 

■ Equipment 

• Sieves, 2- or 5-mm mesh size 

• Drying oven, capable of maintaining a temperature of 105 ± 2 °C 

• Muffle furnace, capable of maintaining a temperature of 550 ± 25 °C 
installed under a fume hood 

• Analytical balance, accuracy 0.01 g 

• Porcelain crucibles or bowls 

• Desiccator with an active drying agent 

■ Sample Preparation 

Use field-moist, sieved (< 5 mm) soil or air-dried, sieved (< 2 mm) soil. 
Dry the soil to 105 °C prior to organic matter determination. 



2 Determination of Chemical and Physical Soil Properties 75 

■ Procedure 

1. Determine the dry mass (m s ) of the soil according to Sect. 2.1. 

2. Heat crucibles or bowls in the muffle furnace at 550 ± 25 °C for 20 min, 
cool in a desiccator and determine tare mass (m t ) to 0.1 g. 

3. Weigh 5-20 g (accuracy 0.01 g) of oven-dried (105 °C) soil (see step 1) 
depending on its organic matter content in crucibles or bowls, and place 
them in the cold muffle furnace. 

4. Heat the muffle furnace gradually to 550 ± 25 °C for 2-4 h until mass 
constancy is achieved. 

5. Open the door and cool the muffle furnace down to 100 °C. 

6. Place the crucibles/bowls in the desiccator and cool them to room tem- 
perature (approx. 1 h). 

7. Measure the mass of the filled crucibles/bowls (m c + m t ) twice. The 
difference of each individual measurements from the mean should not 
exceed 5% of the mean. 

■ Calculation 

1. Calculate the loss of mass (Am; g) after ignition at 550 °C using the 
following equation: 

Am = (m s + m t ) - (m c + m t ) = m s - m c (2.25) 

2. The LOI corresponds to the SOM content and can be calculated using 
the following equation: 

Am 
LOI (%) = x 100 (2.26) 

m s 

Am loss of mass of the soil after ignition at 550 °C (g) 

m s mass of the soil dried at 105 °C (g) 

m t mass of the crucibles/bowls ignited to 550 °C (g) 

m c mass of the soil ignited to 550 °C (g) 

■ Notes and Points to Watch 

• Humus-rich samples should be weighed in the crucibles/bowls in a field- 
moist state and dried and heated in the same crucible. In order to avoid 
dusting the organic samples, the crucibles/bowls should be covered with 
a porcelain lid or a metal mesh. 



76 B.-M. Wilke 

• The incineration of the samples should be controlled. The process is 
complete if black particles cannot be found in the sample or if it has 
a light gray to reddish color. 

• Samples which do not show complete incineration should be treated 
with a few drops of saturated ammonium nitrate solution or hydrogen 
peroxide and heated again to 550 °C for 1 h. 

• The LOI is assumed to be equal in most surface soils. Losses of crystalline 
water of clay minerals and gypsum may result in an overestimation 
of SOM contents. The same is true for carbonate-rich soils, because 
decomposition of CaC0 3 , which starts at temperatures of approx. 500 °C. 
Therefore, the method is mainly recommended for sandy and carbonate- 
free soils and peats. Nevertheless, results for clayey soils and soils rich in 
gypsum can be corrected by subtraction of 0.1% SOM per 1% of clayey 
soil and 0.26% SOM per 1% of gypsum-rich soil. 

• The error caused by the destruction of clay minerals may be avoided by 
pre-heating at 430 °C in an N 2 atmosphere. 

• For peat soils the LOI method is advantageous over the carbon determi- 
nation procedures because the carbon content of these materials varies 
between 40 and 100 mass%. 



2.7 

Soil Nutrients: Total Nitrogen 

■ Introduction 

Objectives. Analysis of total N, the C/N ratio, and inorganic N (ammo- 
nium, nitrate) provides an insight into the nitrogen supply to soil mi- 
croflora and plants. The total N content ranges from < 0. 02% (subsoils) 
to > 2. 5% (peats). A-horizons of mineral soils contain 0.06-0.5% N. Ni- 
trogen, phosphorous, and/or potassium deficiency may limit the microbial 
decomposition (mainly cometabolic) of pollutants in soil. Optimum con- 
ditions are achieved at C:N:P ratios of 100:10:2 (Kastner 2001). Therefore, 
the concentrations of these nutrients have to be analyzed and adjusted if 
necessary. Two methods have gained general acceptance for the determi- 
nation of total N in soils, namely the Kjeldahl and the Dumas methods 
(Bremner 1996). The Kjeldahl method is a wet oxidation procedure, the 
Dumas method a dry oxidation (combustion) method. Both methods have 
been standardized (ISO 11261 1995; ISO 13878 1998). 



2 Determination of Chemical and Physical Soil Properties 11 

2.7.1 

Dry Combustion Method ("Elemental Analysis") 

Principle and Theory. The soil is heated in a purified oxygen stream to a tem- 
perature of > 900 °C. Mineral and organic N species are oxidized and/or 
volatilized. Products are oxides of N (NO x ) and molecular N (N 2 ) mainly. 
After transforming into N 2 by reduction on surfaces of metallic copper, 
the N content is measured by means of thermal conductivity detection 
(method adapted from ISO 13878 1998). 

■ Equipment 

• Balance, capable of weighing accurately to 0.1 mg, or microbalance, ca- 
pable of weighing accurately to 0.01 mg 

• Combustion apparatus to determine total N at a temperature > 900 °C, 
including a detector for measuring the nitrogen gas formed 

• Crucibles of various sizes, e.g. 10 or 20 mL, special requirements being 
given in the manual of the apparatus used 

■ Reagents 

• Combustion gas (oxygen), special requirements being given in the in- 
struction manual of the apparatus used 

• Chemicals and/or catalysts for reduction, oxidation, and/or fixing of 
combustion gases that interfere with the analysis 

• Calibration substances, for example acetanilide (C 8 H 9 NO), amino acids 
of known composition, or soil samples with certified N contents, the N 
content of the calibration substance being as similar to the suspected soil 
N content as possible 

■ Sample Preparation 

Soil samples dried in the air, dried in an oven at a temperature not exceeding 
40 °C, or freeze dried (see Chapt. 1) are sieved (2 mm); if a soil mass < 2 g is 
required for the analysis, mill a representative subsample to 0.1-0.15 mm. 

■ Procedure 

1. Calibrate the apparatus as described in the manufacturer's manual. 

2. Weigh out m x g of the air-dried sample or subsample into a crucible. The 
amount for analysis depends on N contents and on the apparatus used. 



78 B.-M. Wilke 

3. Carry out the analysis according to the manufacturer's manual. 

4. Determine the percentage of water content (mass fraction) according to 
the method described in Sect. 2.1. 



■ Calculation 

1. Normally, the primary results (from the apparatus) are given in mg N 
(Xi) or a mass fraction of N (X 2 ), expressed as a percentage of the air- 
dried soil used (mi ). Calculate total N content (w Nt ), in mg/g, on the basis 
the oven-dried soil according to following equations: 

For primary results given in mg of N: 

X x (100 + w) _ 

w Nt = — x — — (2.27) 

m\ 100 

For primary results, given as percent mass fraction of N: 

(100 + w) 

w Nt = X 2 x 10 x (2.28) 

100 

w N t content of N (mg/g oven-dried soil) 

Xi primary result in N (mg) 

X 2 primary result in percentage N (mass fraction) 

mi mass of air-dried soil for analysis (g) 

w percentage of water content (mass fraction) on the basis of oven- 
dried soil (Sect. 2.1) 

2. If oven-dried samples are used, the N content is calculated as follows: 
For primary results given in mg of N: 

WNt - — (2.29) 

m\ 

For primary results given as percent of mass fraction of N: 

w Nt = X 2 x 10 (2.30) 

■ Notes and Points to Watch 

• Today, several automated elemental analyzers for the determination of 
total N are in use (see Bremner 1996). Most of them can be used for 
determination of total N and total C, others also for hydrogen or sulfur. 



2 Determination of Chemical and Physical Soil Properties 79 

• One of the problems linked with the use of elemental analyzers is sample- 
size limitation. This makes it essential to grind soil samples very finely 
in order to get representative subsamples. 

• Soil pores are filled with air, which contains up to 80% N 2 . Nitrogen gas 
can also enter the combustion cell when it is opened for sample exchange. 
Both facts may lead to overestimation of the soil N content. Therefore, 
sufficient purging should be carried out by oxygen gas flow before the 
combustion step. 

2.7.2 

Modified Kjeldahl Method 

Principle. The method (as in ISO 11261 1995) is based on the Kjeldahl 
digestion. Additional reagents are salicylic acid to avoid loss of nitrates, 
and thiosulfate to detect azo-, nitroso- and, nitrocompounds. Instead of 
selenium, titanium dioxide is used as catalyst. 

Theory. The Kjeldahl method generally employed for determination of 
total N involve two steps: (1) digestion of the sample to convert organic N 
into NHj-N and (2) determination of NHj-N in the digest. The digestion is 
usually performed by heating the sample with H 2 S0 4 containing substances 
that promote the oxidation of organic matter and conversion of organic N 
into NHJ-N. For increasing the temperature K 2 S0 4 or Na 2 S0 4 are used. 
Catalysts such as Hg, Cu, Se, or Ti0 2 increase the rate of oxidation of 
organic matter by H 2 S0 4 . A general equation for the digestion process is 
given below: 

Organic-N + H 2 S0 4 -^(NH 4 ) 2 S0 4 + H 2 + C0 2 

+ other matrix by-products. 

For the determination of NHJ -N several methods are possible, namely ion- 
sensitive electrodes, colorimetric analyses, or analyses by steam distillation 
and titration (Forster 1995b). In the distillation method employed by most 
workers, NHJ -N in digests is converted under excess alkali into NH 3 , which 
is collected in boric acid. By this procedure ammonium borate is formed, 
which is titrated back to boric acid with hydrochloric acid according to the 
following equations: 

2NH 3 + H3BO3 -> (NH 4 ) 2 HB0 3 
(NH 4 ) 2 HB0 3 + 2HC1 -> 2NH 4 C1 + H3BO3 



80 B.-M. Wilke 

■ Equipment 

• Digestion flasks or tubes, of nominal volume 50 mL, suitable for the 
digestion stand 

• Digestion stand, glass tubes placed in holes drilled into an aluminum 
block also being suitable 

• Distillation apparatus, preferably of the Parnass- Wagner type 

• Burette, graduated in intervals of 0.05 mL or smaller 

■ Reagents 

• Salicylic acid/sulfuric acid: dissolve 25 g of salicylic acid in 1 L of cone, 
sulfuric acid (p = 1.84 g/cm 3 ). 

• Potassium sulfate catalyst mixture: grind and thoroughly mix 200 g of 
potassium sulfate, 6 g of copper(II) sulfate pentahydrate, and 6 g of tita- 
nium dioxide with the crystal structure of anatase. 

• Sodium thiosulfate pentahydrate: crush the crystals to form a powder 
that passes through a sieve with 0.25-mm mesh size. 

• NaOH solution ( 10 mol/L). 

• Boric acid solution (H3BO3, 20 g/L). 

• Mixed indicator: dissolve 0.1 g of bromocresol green and 0.02 g of methyl 
red in 100 mL of ethanol. 

• Sulfuric acid (0.01 mol/L). 

■ Sample Preparation 

Soil samples dried in the air, dried in an oven at temperature not exceeding 
40 °C, or freeze-dried (see Chapt. 1) are sieved (2 mm) and ground to 
0.1-0.15 mm. 

■ Procedure 

1. Place a test portion of the air-dried soil sample of about 0.2 g (expected 
N content ca. 0.5%) to 1 g (expected N content ca. 0.1%) in the digestion 
flask. 



2 Determination of Chemical and Physical Soil Properties 81 

2. Add 4 mL of salicylic/sulfuric acid and swirl the flask until the acid is 
thoroughly mixed with the soil. 

3. Allow the mixture to stand for at least several hours (or overnight). 

4. Add 0.5 g of sodium thiosulfate through a dry funnel with a long stem 
that reaches down into the bulb of the digestion flask, and heat the 
mixture cautiously on the digestion stand until frothing has ceased. 

5. Cool the flask. 

6. Add 1.1 g of the catalyst mixture and heat until the digestion mixture 
becomes clear. 

7. Boil the mixture gently for up to 5 h so that the sulfuric acid condenses 
about 1/3 of the way up to the neck of the flask. Ensure that the tem- 
perature of the solution does not exceed 400 °C. In most cases a boiling 
period of 2 h is sufficient. 

8. After completion of the digestion step, allow the flask to cool, and add 
about 20 mL of water slowly while shaking. 

9. Swirl the flask to bring any insoluble material into suspension and 
transfer the contents to the distillation apparatus. 

10. Rinse three times with water to complete the transfer. 

11. Add 5 mL of boric acid solution to a 100 mL conical flask and place the 
flask under the condenser of the distillation apparatus in such a way 
that the end of the condenser dips into the solution. 

12. Add 20 mL of sodium hydroxide solution to the funnel of the apparatus 
and run the alkali slowly into the distillation chamber. 

13. Distill about 40 mL of condensate (the amount depends on the dimen- 
sions of the conical flask). 

14. Rinse the end of the condenser. 

15. Add a few drops of indicator to the distillate. 

16. Titrate with sulfuric acid to a violet endpoint. 

■ Calculation 

Calculate total N content (w Nt ) using the following equation and round the 
result to two significant figures: 

< xt/ -n (^i " ^o) x c(H + ) x M N 100 + w H2 o „ al , 

w Nt (mg N/g soil) = x — (2.31) 

m 100 



82 B.-M. Wilke 

V\ volume of the sulfuric acid (mL) 

Vq volume of the sulfuric acid used in the blank test (mL) 

c(H + ) concentration of H + in the sulfuric acid (mol/L; e.g., if O.Olmol/L 
sulfuric acid is used, c(H + ) is 0.02 mol/L 

M N molar mass of nitrogen (g/mol; 14) 

m mass of the air-dried soil subsample used (g) 

Wh 2 o water content expressed as a percentage by mass on the basis of 
oven-dried soil (Sect. 2.1) 

■ Notes and Points to Watch 

• The modified Kjeldahl procedure is satisfying for the analyses of most N 
compounds in soils, but it detects compounds containing N-N and N-O 
linkages and some heterocyclics (e.g., pyridine) only partially (Bremner 
1996; ISO 11261 1995). 

• Losses of nitrogen can occur with samples of high NH 4 -N and NO3-N 
content. Therefore, excessive drying prior to analysis should be avoided. 

• A potentiometric titration is also possible. The endpoint of the titration 
is pH 5.0. 

• If a steam distillation is used, a distillation rate up to 25 mL/min is 
applicable. Stop the distillation when about 100 mL of distillate have 
been collected. 

2.8 

Soil Nutrients: Inorganic Nitrogen 

■ Introduction 

Objectives. Nitrogen is a main nutrient for plants and soil organisms. Am- 
monium and nitrate in the soil are the N sources immediately available 
to plants. They are produced by mineralization of organic compounds or 
fertilized to soil. Besides ammonium and nitrate, nitrite (N0 2 ) may also 
be present, although its content is usually negligible except in neutral and 
alkaline soils recently treated with NHJ salts or NHJ -forming fertiliz- 
ers (Mulvaney 1996). Quantification of nitrate and ammonium N in soil 
extracts and soil solution can be performed using colorimetric, microdif- 
fusion, and ion electrode methods (Mulvaney 1996). 

Principle. Fresh soil is extracted with a calcium chloride solution (0.01 mol/ 
L) in a 1:10 ratio (m/v) at 20 ± 1 °C. After reaching equilibrium (2 h), the 



2 Determination of Chemical and Physical Soil Properties 83 

solution is centrifuged for the determination of ammonium and nitrate. 
A segmented flow analysis (SFA) system equipped with a colorimetric 
detector is used to quantify nitrate and ammonium N. 

Theory. Up to 90% of total N in upper soil layers exists in organic forms. Or- 
ganic N is mineralized by soil microorganisms to ammonium, nitrite, and 
nitrate, which are easily available to plants. In loamy and clayey soils 20- 
25% of total N are present as fixed ammonium in clay minerals (Li et al. 1990; 
Sahrawat 1995). Fixed ammonium occurs between the layers of 2: 1 -type clay 
minerals. It cannot be replaced by neutral potassium solution (Mulvaney 
1996). In contrast to exchangeable ammonium, it is not available to plants. 
The method described here is adapted from ISO 14255 (1998). It is valid 
for the determination of soluble inorganic N and weakly adsorbed am- 
monium. In acid clayey forest soils, ammonium specifically bound at the 
edges of clay mineral interlayers may be the main N source of plants. For 
the determination of available N in these soils the use of KC1 (1 mol/L) is 
recommended as extractant (VDLUFA 2000). 

2.8.1 
Extraction 

■ Equipment 

• Balance, accuracy 10 mg 

• Polyethylene bottles, nominal volume 250 mL, with screwcaps 

• Shaking machine or reciprocating shaker, 150-250 rpm 

• Centrifuge, capable of holding the tubes used 

• Polyethylene centrifuge tubes, nominal volume 100 mL or other sufficient 
volume 

■ Reagents 

• Water, with a specific conductivity not higher than 0.2mS/m at 25 °C 
(according to grade 2 of ISO 3696 1987) 

• Extraction solution: CaCl 2 (0.01 mol/L) 

■ Sample Preparation 

Field-moist soil samples shall be stored in a cooling box immediately after 
sampling. In the laboratory they are homogenized, sieved (< 2 or < 5 mm), 
and analyzed immediately. If the analysis cannot be carried out the same 
day, storing at 4°C (up to 5 days) or freezing at -25 °C is necessary. In 



84 B.-M. Wilke 

order to avoid microbial transformation of soil N, slow thawing of frozen 
samples should be avoided. Homogenized and frozen samples stored in 
plastic bags can be reduced to small pieces by slamming the frozen plastic 
bags on a hard pad. A separate part of the homogenized sample is used for 
the determination of the water content (see Sect. 2.1). 

■ Procedure 

1. Weigh 10.00 g of the laboratory sample in a polyethylene bottle. 

2. Add 100 mL of extraction solution at a temperature of 20 °C and shake 
mechanically for 2 h. Perform a blank test by adding only the extraction 
solution to the polyethylene bottle. 

3. Decant the required quantity of the extract suspension into centrifuge 
tubes and centrifuge for 10 min at about 3,000 g. 

4. Decant the supernatant solution in measuring cups and measure the 
contents of nitrate, nitrite (if necessary), and ammonium as described 
below (see Sects. 2.8.2, 2.8.3). 

■ Notes and Points to Watch 

• In order to avoid microbial transformation of N, sampling and storage 
conditions described above have to be observed strictly. For the handling 
of large sample series, quick drying at 105 °C or air drying is used. In both 
cases microbial transformations cannot be avoided totally. Furthermore, 
drying at high temperatures increases the ammonium content of soil 
samples significantly (VDLUFA 2000). 

• Sampling of arable soils should be carried out in early spring. At this 
time inorganic N mainly exists as nitrate. 

• If the homogenization of samples is difficult, larger sample weights (up 
to 150 g) are recommended (VDLUFA 2000). In any case the soil solution 
ratio shall be kept at 1:10 (m/v). 

• The soluble N fractions should be measured immediately if possible, but 
not later than 1 day after the extraction. If this is not possible, the extracts 
should be stored in a refrigerator at a temperature not exceeding 4 °C for 
a maximum of 1 week. 

2.8.2 

Quantification of Nitrate Nitrogen 

Principle. In an SFA system, the sample is first subjected to dialysis. Nitrate 
and nitrite ions of the samples pass through the membrane. The nitrate 



2 Determination of Chemical and Physical Soil Properties 85 

is then reduced to nitrite by means of cadmium. Next a-naphthylethyl- 
enediamine dihydro chloride and sulfanilamide are added, so that a red- 
colored diazo compound is formed in the acidic medium. Its absorbance is 
measured at a wavelength of 543 nm. 

■ Equipment 

• SFA system consisting of a sampler, pump, dialysis unit, reduction col- 
umn, nitrate unit, photometer, and recorder 

• Cd/Cu reduction tube consisting of U-shaped glass tubing about 15 cm 
long, with internal diameter of 2 mm, and provided with ferrule for 
connection to the SFA tubing (may be purchased from the SFA system 
manufacturer) 

■ Reagents 

• Wetting agent, polyoxyethylene lauryl ether (30%). 

• Buffer solution: dissolve 25 g of NH 4 C1 in water, add 12.5 mL of NH 4 OH 
solution (3%) and 1 mL of wetting agent. Make up to 1 L with water and 
mix. 

• Cd/Cu reducing agent: swirl approx. 5 g of cadmium powder (particle 
size 0.3-0.8 mm) for 1 min with about 30 mL of HC1 ( 1 mol/L). Wash with 
water until acid free. Then add about 50 mL of a CuS0 4 solution (20 g/L) 
and swirl for 3 min. Wash at least ten times with water to remove any 
flocculated copper. Store the Cd/Cu reducing agent in a dark place. 

• Cd/Cu reduction tube: fill the U-shaped column with buffer solution, 
taking care not to introduce air bubbles. Introduce the activated cad- 
mium powder with the aid of a funnel on both sides of the column. 
Apply vibration now and then to pack the powder. Fill the column up to 
5 mm from the top and seal the ends with small plugs of glass wool. The 
column is now ready for use and can be placed in the SFA system. 

• Color reagent: in a 1 L volumetric flask, add 150 mL of cone. H3PO4 
(85%) to 0.5 L of water. Add 0.5 g of a-naphthylethylenediamine dihy- 
drochloride (Ci 2 H 16 N 2 Cl2) and swirl until dissolved. Then dissolve 10 g 
of sulfanilamide (C6H 8 N 2 S) in this mixture and fill up to the mark with 
water. 

■ Procedure 

The analysis of nitrate nitrogen in calcium chloride soil extracts is carried 
out with the SFA system. Details are given in the manufacturer's manual. 



86 B.-M. Wilke 

2.8.3 

Quantification of Ammonium Nitrogen 

Principle. In an SFA system, the sample is first subjected to dialysis. The 
determination of ammonium is based on the Berthelot reaction, in which 
a phenol derivative (here salicylate) forms an indophenol in the presence of 
ammonia and hypochlorite under the catalytic action of sodium nitroferri- 
cyanide (nitroprusside). In alkaline medium, the indophenol thus formed 
has a green-blue color, the absorbance of which is measured at a wavelength 
of 660 nm. 

■ Reagents 

• Buffer solution, pH 5.2: dissolve 24 g of sodium citrate (Na 3 C 6 H 5 07) and 
33 g of sodium potassium tartrate (KNaC 4 H 4 6 ) and make up to 1 L with 
water. Add 1 mL of wetting agent (30% polyoxyethylene lauryl ether. 

• Color reagent: in a 1 L volumetric flask containing about 800 mL of water, 
dissolve 80 g of sodium salicylate (C 7 H 5 3 Na) and 25 g of NaOH. Fill up 
to the mark with water. 

• Nitroferricyanide (nitroprusside) solution: dissolve 1 g of sodium nitro- 
ferricyanide dihydrate (Na 2 [Fe(CN) 5 NO] x 2H 2 0) in 1 L of water. 

• Isocyanurate solution: Dissolve 2g of sodium dichloroisocyanurate 
(Cl2C 3 N 3 Na03 x 2H 2 0) and 25 g of NaOH in 1 L of water. 

■ Procedure 

The analysis of ammonium nitrogen in calcium chloride soil extracts is 
carried out with the SFA system. Details are given in the manufacturer's 
manual. 

■ Calculation 

The content of the different N fractions in the soil material (w N ), expressed 
in mg/kg, is calculated using the following equation: 

(a-b)x 10x(100 + w) 

w N = (2.32) 

100 

a content of NO3-N and NH 4 -N in the soil extract (mg/L) 

b content of NO3-N and NH 4 -N in the blank extract (mg/L) 

w percentage of water content (m/m) on the basis of the air-dried soil 
(Sect. 2.1) 



2 Determination of Chemical and Physical Soil Properties 87 

2.9 

Soil Nutrients: Phosphorus 

■ Introduction 

Objectives. Soil phosphorus is, besides nitrogen, potassium, calcium, and 
magnesium, a main nutrient for soil organisms and plants. It exists in 
inorganic and organic fractions with varying percentages between 5 and 
95%. The soil organic P fraction may be derived from plant residues, soil 
flora, and soil fauna tissues and residues that resist rapid hydrolysis (Kuo 
1996). Inorganic fractions consist of Ca-, A1-, and Fe-phosphates. The most 
prominent phosphate mineral in soils is apatite (Ca 5 (P0 4 ) 3 OH). The to- 
tal concentration in soil is generally in the range from 200 to 800 mg/kg. 
A considerable amount of P is also bound in the amorphous mineral frac- 
tion. Part of this is specifically adsorbed on surfaces of iron and aluminum 
oxides. Only a small part appears in a soil solution (< 0.1 mg/L in unfertil- 
ized soils and subsoils, 0.1-5 mg/L in Ap-horizons of arable soils (Scheffer 
2002). 

Soil microbes are involved in the mineralization of P from organic debris 
(Forster 1995c). Extracellular phosphatases are produced by microorgan- 
isms and roots and contribute to the mineralization of organic P. Deficiency 
of P may limit the growth of plants and the microbial decomposition of pol- 
lutants in soil. P is likely to be deficient in hydrocarbon-impacted soils and 
subsoils. Therefore, its concentration has to be analyzed and, if necessary, 
adjusted. Regarding available fractions of nutrients, optimum conditions 
are achieved at C:N:P ratios of 100:10:2 (Kastner 2001). 

There are several methods available for the determination of total, in- 
organic, organic, and labile (plant available) P (VDLUFA 1991; Kuo 1996). 
Only a minor amount of soil P is available to plants. The estimation of plant 
available P is essential for optimum nutrient supply and can be applied 
within the scope of phytoremediation. Numerous tests have been developed 
to extract varying amounts of P, depending on the types of extractants used 
(e.g., CaCl 2 , lactate, acetate, EDTA, ammonium bicarbonate; see also Kuo 
1996). The test described here uses a sodium hydrogen carbonate solution. 
It is useful for both acid and calcareous soils and has been standardized 
(ISO 11263 1994). There are several methods for the quantification of P in 
soil extracts and soil solution, namely, spectophotometry (most common), 
ion chromatography, and inductively coupled plasma spectrometry (for 
details see Kuo 1996). The spectrophotometric method described below 
was developed by Murphy and Riley (1962). 

Principle. Total phosphorus is extracted from finely ground soil with cone, 
sulfuric acid, hydrogen peroxide, and hydrofluoric acid (Bowman 1988). 
Labile P is extracted with a sodium hydrogen carbonate solution at pH 8.50. 



88 B.-M. Wilke 

Clear extracts are quantitatively analyzed for P by a spectrophotometric 
method involving the formation of an antimony-phosphate-molybdate 
complex reduced with ascorbic acid to form a deep-blue-colored complex. 

Theory. Organic and non-silicate inorganic forms of P are dissolved in 
sulfuric acid and hydrogen peroxide. P in silicate lattice is released by the 
hydrofluoric acid treatment. Labile P is extracted using a NaHC0 3 solution. 
The OH" and CO3" ions in the NaHC0 3 solution decrease the concentration 
or activity of Ca 2+ and Al 3+ in the soil solution, resulting in an increased 
solubility of P. Quantification of P is based on the reaction of phosphoric 
acid with molybdate ions, which forms a heteropoly molybdophosphate 
complex: 

H3PO4 + 12H 2 Mo0 4 -> H 3 P(Mo 3 Oio)4 + 12H 2 

The complex has a yellow color. In the presence of reducing agent such as 
ascorbic acid, the Mo in the complex is partially reduced from 6+ to 3+ 
and/or 5+, which results in a characteristic blue color. 



2.9.1 

Extraction of Total Phosphorus 

■ Equipment 

• Pebble mill for grinding sieved soil 

• Analytical balance, accuracy 0.01 mg 

• Hot plates or heated sand bath 

• Teflon beakers, 100 mL 

■ Reagents 

• Cone, sulfuric acid (95-97%) 

• Hydrogen peroxide (30%) 

• Cone, hydrofluoric acid (40%) 

■ Sample Preparation 

Air-dried soil samples are sieved (2 mm) and ground to 0.1-0.15 mm using 
a pebble mill. The dry mass portion (percentage) is determined as described 
in Sect. 2.1. 



2 Determination of Chemical and Physical Soil Properties 89 

■ Procedure 

1. Weigh 0.5 g finely ground, well mixed soil into a 100 mL Teflon beaker. 
For high organic matter soils use 0.25 g. 

2. Add 5 mL of cone, sulfuric acid and swirl gently. 

3. Add 3 mL of 30% hydrogen peroxide in 0.5 mL portions. 

4. Swirl vigorously (caution: foaming may lead to an overflow of samples 
high in organic matter). 

5. When the reaction with hydrogen peroxide has subsided, add 1 mL of 
cone, hydrofluoric acid in 0.5 mL portions and swirl gently. 

6. Place the beaker on a hot plate at 150 °C for 10-20 min to eliminate 
excess hydrogen peroxide. 

7. After slight cooling, wash down the sides of the beaker with approx. 
15 mL distilled water. 

8. Mix and cool to room temperature. 

9. Transfer the beakers content quantitatively to a 50 mL volumetric flask, 
passing it through a filter paper. 

10. Make two additional washings of the beaker with 10 mL of distilled 
water, filter, and make up to volume. 

11. Measure the P concentration of the extract using the molybdenum blue 
method described in Sect. 2.9.3. 

■ Calculation 

The total content of P (w P t; mg/kg oven-dried soil) is calculated using the 
following equation: 

p P x50 (100 + w w ) ,___. 

WpT = x (2.33) 

m 100 

Pp concentration of P (mg/L) measured according to the method de- 
scribed in Sect. 2.9.3 

m mass of air-dried soil (g) 

ww percentage of water content (mass fraction) on the basis of oven-dried 
soil (Sect. 2.1) 



90 B.-M. Wilke 

■ Notes and Points to Watch 

• The extraction procedure described may be also used for determination 
of total contents of other elements (e.g., K, Ca, Mg, Al, Fe). 

2.9.2 

Extraction of Labile Phosphorus 

■ Equipment 

• Analytical balance, with a readability of ±0.01 g 

• Shaker, end over end (30-35 rpm) or eccentric horizontal (140 thrusts 
per min) 

• pH meter, with a readability of ±0. 01 pH units 

• Fluted filters, free of P, fine to medium porosity 

• Apparatus to filter extracts simultaneously 

■ Reagents 

• Sodium hydroxide solution (NaOH, 1 mol/L); store in an inert hermeti- 
cally sealed bottle. 

• Extracting solution: dissolve 42 ± 0.1 g of sodium hydrogen carbonate 
(NaHC0 3 ) in 800 mL of water. Adjust the pH to 8. 50 ± 0. 02 with sodium 
hydroxide solution. Make up the solution to 1 L with water. 

• Carbon, activated, allowing the absorbance of the blank to be less than 
0.015. 

■ Sample Preparation 

Soils samples are dried in the air, or in an oven at temperature not exceeding 
40 °C, or freeze-dried (see Chapt. 1). Then samples are sieved (2 mm). The 
dry mass portion (percentage) is determined according to Sect. 2.1 

■ Procedure 

1. Weigh 5.00 ± 0.01 g of pretreated soil into a 250-mL flask. 

2. Add 1.0 g of activated carbon and 100 ± 0.5 mL of extracting solution. 

3. Stopper the flask and place it immediately on the shaker. 



2 Determination of Chemical and Physical Soil Properties 91 

4. Shake for exactly 30 min at 20 ± 1 °C. 

5. Filter immediately (within 1 min) into a dry vessel using a P free-fluted 
filter paper. 

6. Prepare a blank by following the above procedure without soil. 

7. Measure the P concentration of filtrates as given below. 

■ Calculation 

The content of phosphorous soluble in sodium hydrogen carbonate (w p ; 
mg/kg of oven-dried soil) is calculated using the following equation: 

100 + w w , 

Wp = Pp x 20 x (2.34) 

Y 100 

p P concentration of P (mg/L) as measured in the extract according to the 
method described in Sect. 2.9.3 

20 quotient of the volume of extracting solution (100 mL) and the mass 
of air-dried soil (5 g) 

w w percentage of water content (mass fraction) on the basis of oven-dried 
soil (Sect. 2.1) 

■ Notes and Points to Watch 

• The extracting solution must be used within 4 h after preparation. 

• A NaHC0 3 test level of 10 mg P/kg soil is considered to be in the "high" 
category (see Kuo 1996). 

2.9.3 

Quantification of Phosphorus 

■ Equipment 

• Analytical balance, with a readability of ±0.001 g 

• Spectrophotometer, capable of measuring the absorbance in wavelength 
up to 900 nm and accepting cells of path 10 mm (readability 0.001 units 
of absorbance) 

• Optical cells, of path length 10 mm 

• Volumetric flasks, 50 mL 



92 B.-M. Wilke 

■ Reagents 

• Sulfuric acid, (H 2 S0 4 2.5 mol/L) 

• Ammonium molybdate solution: dissolve 20 g of ammonium hepta- 
molybdate tetrahydrate ([(NH 4 ) 6 Mo 7 2 4 4H 2 0]) in 500 mL of deionized 
water. Store the solution in a glass-stoppered bottle. 

• Antimony potassium tartrate solution (1 mg Sb/mL): dissolve 0.2728 g of 
K(SbO) C 4 H 4 6 x 1/2H 2 in 100 mL of deionized water. 

• Ascorbic acid solution (0.1 mol/L): dissolve 1.76 g of C6H 8 06 in 100 mL 
of deionized water. Prepare the solution fresh daily. 

• Mixed reagent: thoroughly mix 50 mL of H 2 S0 4 (2.5 mol/L), 15 mL of 
ammonium molybdate solution, 30 mL of ascorbic acid solution, and 
5 mL of antimony potassium tartrate solution. Prepare fresh daily. 

• Phosphate stock solution (50mgP/L): dissolve 0.2197 g of oven-dried 
(40 °C) KH 2 P0 4 in deionized water. Add 25 mL of H 2 S0 4 (2.5 mol/L) and 
dilute to 1 L with deionized water. 

• Working phosphate standard solution (5mgP/L): dilute 10 mL stock 
solution to 100 mL with deionized water. 

■ Sample Preparation 

Soil samples are extracted as described in Sects. 2.9.1 and 2.9.2. 

■ Procedure 

1. Transfer into 50 mL volumetric flasks: 

- A standard series, ranging from 0.5 mL up to 8mL of the working 
phosphate standard solution, corresponding to P concentrations in 
the measuring solution between 0.05 and 0.8 mg/L. 

- An aliquot of lOmL of the soil extract containing total phosphorus 
if the expected concentration of P in the soil is less than 400 mg/kg; 
otherwise use 5 mL or less. 

- An aliquot of 25 mL of the soil extract containing labile phosphorus. 

2. Dilute the aliquots with deionized water to about 25 mL (if necessary), 
and add 8 mL of mixed reagent. 

3. Dilute the solution to volume and mix well. 

4. Measure the absorbance at 880 nm after 10 min in a spectrophotometer. 

5. Prepare a blank that contains all reagents except the P solution. 



2 Determination of Chemical and Physical Soil Properties 93 

■ Calculation 

For the evaluation of the spectrophotometric measurements prepare a cal- 
ibration graph by plotting absorbance units versus the P concentrations of 
standard solutions (mg P/L). The correlation between both parameters is 
linear in the relevant range up to 0.8 mg P/L. 

The phosphorous concentration of the measuring solution (p P ; mg/L) 
can be calculated using the following equation: 

(A ES - A B ) x 50 
Pp = 7 77 ( 2 - 35 ) 

/x y s 

A E s absorbance of the soil extract 

A B absorbance of the blank 

/ slope of the regression line (absorbance per mg P/L) 

V s volume of the aliquot (mL) 

50 volume of the volumetric flasks (mL) 

References 

Blake JR, Hartge KH (1986) Bulk density. In: Klute A (ed) Methods in soil analysis, part 1. 

Physical and mineralogical methods, 2nd ed, Am Soc Agron, Madison, WI, pp 363-376 
Bowman RA (1988) A rapid method to determine phosphorus in soils. Soil Sci Soc Am J 

52:1301-1304 
Bremner JM (1996) Nitrogen - total. In: Bigham JM (ed) Methods of soil analysis, part 3, 

chemical methods. Soil Sci Soc Am Am Soc Agron, SSSA Book, Series no 5, Madison, 

WI,pp 1085-1121 
Danielson RE, Sutherland PL (1986) Porosity. In: Klute A (ed) Methods in soil analysis, 

part 1, physical and mineralogical methods, 2nd edn. Soil Sci Soc Am Am Soc Agron, 

Madison WI,pp 443-461 
DIN 19684-3 (1977) Bodenuntersuchungsverfahren fur den Landwirschaftlichen Wasser- 

bau - Chemische Laboruntersuchungen - Teil 3: Bestimmung des Gliihverlustes und 

des Gliihriickstandes 
Forster JC (1995a) Soil sampling, handling, storage and analysis - organic carbon - In: 

Alef K, Nannipieri P (eds) Methods in applied soil microbiology and biochemistry. 

Academic Press, pp 59-65 
Forster JC (1995b) Soil sampling, handling, storage and analysis - soil nitrogen. In: Alef K, 

Nannipieri P (eds) Methods in applied soil microbiology and biochemistry. Academic 

Press, pp 79-87 
Forster JC (1995c) Soil sampling, handling, storage and analysis - soil phosphorus. In: 

Alef K, Nannipieri P (eds) Methods in applied soil microbiology and biochemistry. 

Academic Press, pp 88-93 
Hartge KH (1968) Heterogenitat des Bodens oder Quellung? Trans Int Congr Soil Sci 3:591- 

597 



94 B.-M. Wilke 

ISO 10693 (1995) Soil Quality - Determination of carbonate content - volumetric method 

ISO 10694 (1995) Soil Quality - Determination of organic and total carbon after dry com- 
bustion (elementary analysis) 

ISO 11261 (1995) Soil Quality- Determination of total nitrogen- Modified Kjeldahl method 

ISO 11263 (1994) Soil Quality - Determination of phosphorous - spectrometric determina- 
tion of phosphorous soluble in sodium hydrogen carbonate solution 

ISO 1 1266 ( 1 994) Soil Quality - Guidance on laboratory testing for biodegradation of organic 
chemicals in soil under aerobic conditions 

ISO 11272 (1998) Soil Quality - Determination of dry bulk density 

ISO 11274 (1998) Soil Quality - Determination of the water- retention characteristic - Lab- 
oratory methods 

ISO 11465 (1993) Soil Quality - Determination of dry matter and water content on a mass 
basis - Gravimetric method 

ISO 13878 (1998) Soil Quality - Determination of total nitrogen content by dry combustion 
("elemental analysis") 

ISO 14238 (1997) Soil quality - Biological methods - Determination of nitrogen mineral- 
ization and nitrification in soils and the influence of chemicals on these processes 

ISO 14255 (1998) Soil Quality - Determination of nitrate nitrogen, ammonium nitrogen 
and total soluble nitrogen in air-dry soils using calcium chloride solution as extractant 

ISO 16072 (2002) Soil quality - Laboratory methods for determination of microbial soil 
respiration 

ISO 3696 (1987) Water for analytical laboratory use - Specifications and test methods 

ISO DIS 10390 (2002) Soil Quality - Determination of pH 

ISO/TS 14256-1 (2003) Soil Quality - Determination of nitrate, nitrite and ammonium in 
field moist soils by extraction with potassium chloride solution - Part 1 : Manual method 

Kastner M (2001) Parameter und Methoden zur Beurteilung der biologischen Sanierbarkeit 
von Boden. In: Michels J, Track Th, Gehrke U, Sell D (eds) Leitfaden, Biologische 
Verfahren zur Bodensanierung. DECHEMA Gesellschaft fur Chemische Technik und 
Biotechnologie eV, Bericht im Auftrag des BMBF Projekttrager Abfallwirtschaft und 
Altlastensanierung im Umweltbundesamt Berlin. Forderkennzeichen 1491064, pp 191 — 
234 

Kuo S (1996) Phosphorous. In: Bigham JM (ed) Methods of soil analysis, part 3, chemical 
methods. Soil Sci Soc Am Am Soc Agron, SSSA Book, Series no 5, Madison WI, pp 869- 
919 

In: Bigham JM (ed) Methods of soil analysis, part 3, chemical methods. Soil Sci Soc Am Am 
Soc Agron, SSSA Book, Series no 5, Madison WI, pp 1123-1184 

Li C, Fan X, Mengel K (1990) Turnover of interlayer ammonium in loess-derived soil grown 
with winter wheat in the Shaanxi Province of China. Biol Fert Soils 9:21 1-214 

Mulvaney RL (1996) Nitrogen - inorganic forms. In: Bigham JM (ed) Methods of soil 
analysis, part 3, chemical methods. Soil Sci Soc Am Am Soc Agron, SSSA Book, Series 
no 5, Madison WI, pp 1123-1184 

Murphy J, Riley HP (1962) A modified single solution method for the determination of 
phosphate in natural waters. Anal Chim Acta 27:31-36 

Nelson, DW, Sommer LE (1996) Total carbon, organic carbon, and organic matter. In: 
Bigham JM (ed) Methods of soil analysis, part 3, chemical methods. Soil Sci Soc Am 
Am Soc Agron, SSSA Book, Series no 5, Madison WI, pp 961-1010 

Page AL, Miller RH, Keeney DR (eds, 1982) Methods in soil analysis, part 2, chemical and 
microbiological properties. Soil Sci Soc Am Am Soc Agron, SSSA Book, Madison WI 

Sahrawat KL ( 1 995) Fixed ammonium and carbon-nitrogen ratios of some semi-arid tropical 
Indian soils. Geoderma 68:219-224 



2 Determination of Chemical and Physical Soil Properties 95 

Scheffer F (2002) Lehrbuch der Bodenkunde, Scheffer/Schachtschabel. 15. Auflage, neubear- 
beitet und erweitert von Blume HP et al. Akademischer Verlag GmbH, Heidelberg 

Schlichting E, Blume H-P, Stahr K (1995) Bodenkundliches Praktikum, 2nd ed, Blackwell, 
Berlin Wien 

Topp GC, Dow B, Edwards M, Gregorich EG, Curnoe WE, Cook EJ (2000) Oxygen measure- 
ments in the root zone facilitated by TRD. Can J Soil Sci 80:33-41 

VDLUFA (2000) Methodenbuch 1 3. Teilliefg. 2000, Nmin-Methode Labor A6. 1.4.1, VDLUFA- 
Verlag Darmstadt 



3 



Quantification of Soil Contamination 

Kirsten S. j0rgensen, Olli Jarvinen, Pirjo Sainio, 
Jani Salminen, Anna-Mari Suortti 



3.1 

General Introduction 

Even though better toxicity tests have become available and their use is 
increasing in risk assessment of contaminated sites and of the reuse of 
bioremediated soil, chemical data is still today the main information used 
in decision-making. The awareness of soil contamination increased in the 
1980s and large-scale bioremediation became frequent during the 1990s. 
However, the development and standardization of reproducible chemical 
methods for the determination of specific organic contaminants in soils 
have been very slow (Karstensen et al. 1998). The methods for the de- 
termination of heavy metals have been in use longer, metal examination 
of soil samples having been performed for geological purposes, e.g., in 
the search for ore by the mining industry. The older methods for deter- 
mination of oil and polyaromatic hydrocarbons (PAHs) were often based 
on extraction of dried and sieved samples (resembling the procedures for 
heavy metal pretreatment) followed by an extraction with non-polar sol- 
vents. The use of halogenated solvents such as CC1 4 and Freon (ISO/TR 
11046 1994) were common, but due to occupational health and environ- 
mental aspects these are being phased out. Methods that are based on the 
extraction of field-moist samples with a mixture of polar (e.g., acetone, 
methanol) and non-polar (hexane, pentane, heptane) solvents have been 
proven to give sufficient yield and to be reproducible. Methods that are fea- 
sible and reproducible in the laboratory are currently being standardized 
by the International Standardization Organization (ISO). 

The objectives of determining contaminant concentrations in soil may 
be to assess the appearance of particular contaminants at a site or to 
monitor the progress of a bioremediation action either in the field or in 
laboratory feasibility studies. The monitoring of bioremediation involves 
repeated measurements of contaminant concentrations over time. Based on 
a time series with, e.g., five points, a biodegradation rate can be obtained. 



Kirsten j0rgensen, Olli Jarvinen, Pirjo Sainio, Jani Salminen: Finnish Environment Institute, 
P.O. Box 140, 00251 Helsinki, Finland, E-mail: Kirsten.Jorgensen@ymparisto.fi 

Anna-Mari Suortti: SGS Inspection Services, Syvasatamantie 24, 49460 Hamina, Finland 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



98 K.S. j0rgensen et al. 

The biodegradation rate can be linear or represent first order decay depen- 
dent on, e.g., the contaminant concentration and bioavailability. Field-scale 
bioremediation can be classified as either in situ methods, where the treat- 
ment takes place without excavating the soil, and ex situ methods, where ex- 
cavated soil is treated typically in piles. When monitoring a site undergoing 
in situ treatment by drilling for subsurface samples, it is essential to remem- 
ber that true replicate samples cannot be obtained and a large variation is to 
be expected. When sampling stock piles or biopiles, a combination sample 
consisting of subsamples from different places in the piles typically will be 
assembled, and parallel combination samples can be made ( j0rgensen et al. 
2000). When monitoring biodegradation by laboratory microcosms, it is of 
great importance that all the sample material representing a certain depth, 
treatment, etc., is homogenous. This is best ensured by homogenizing and 
sieving a larger batch from the field and by distributing this into separate 
parallel bottles or other containers for laboratory incubations. A mesh size 
of 8 mm has proven to be a good size for sieving field-moist soil (Laine 
and j0rgensen 1997; Salminen et al. 2004). However, the measurement of 
contaminant disappearance only shows that the parent compound has been 
transformed; it does not reveal whether the degradation is complete to C0 2 
or CH 4 or if other degradation products are produced. 

Contamination with petroleum hydrocarbon products is one of the most 
frequent types of soil contamination. Refineries, surface and underground 
storage tanks, petrol service stations, etc., are the most common sites for 
such contamination. Most petroleum products also contain minor amounts 
of PAHs. No single method is reliable for the determination of all petroleum 
hydrocarbons, and we therefore describe three methods for the determi- 
nation of different fractions of hydrocarbons in soil samples. 

Volatile hydrocarbons (Sect. 3.2) should be determined at sites where 
gasoline and jet fuel are the sources of contamination. The pertinent 
method here quantitatively determines these separate compounds: ben- 
zene, toluene, ethylbenzene and xylenes (BTEX compounds), naphthalene, 
and gasoline additives such as MTBE (methyl tert-butyl ether) and TAME 
(tert-amyl methyl ether). This method can also be used to determine halo- 
genated volatiles, which may often be found together with fuel products 
because such solvents often are used, e.g., for cleaning engines. 

Contamination with oil products such as heating oil, diesel or lubricating 
oil is best determined using the method (Sect. 3.3) for hydrocarbons in 
the range C 10 to C 40 . The result is a sum parameter, which does not give 
concentrations of specific compounds. But still the sum of the hundreds of 
compounds in this range is very useful for quantifying contamination with 
them and for monitoring bioremediation. Based on the chromatogram, 
a qualitative estimation of the type of contamination can be obtained. 
This Qo-40 parameter is often referred to as mineral oil or total petroleum 



3 Quantification of Soil Contamination 99 

hydrocarbons (TPH), but these terms are somewhat unspecific. Crude oil is 
often determined with this method, but it also includes volatiles and PAHs 
that should be determined separately with the methods for volatiles and 
PAHs, respectively. 

Contamination with PAHs is commonly found at gas works and at sites 
where coal tar and oil shale are handled. Oil containing heavy fractions 
or waste oil may also contain significant amounts of PAHs. The method 
described here (Sect. 3.4) allows for a single determination of 16 different 
PAH compounds. In the literature the sum of PAHs is often reported, but 
the fact that different countries and different laboratories analyze different 
number of compounds has made this term very unspecific. Guideline values 
for clean-up needs also differ between countries, so it is important to 
check which compounds require reports. Since the toxicities of the PAH 
compounds differ, there may not be any guideline value set out for all 
compounds. 

Contamination with heavy metals is difficult to assess because clean 
soil itself may contain many heavy metals, depending on the geological 
structure. Furthermore, many metals are not necessarily bioavailable in 
soil, and for that reason different types of less exhaustive extractions are 
being developed to determine the bioavailable fractions. The background 
contents of metals in soil are in many countries known and they are taken 
into account when guideline values for clean-up are determined. Still today 
most guideline values are based on the total or near- total content of metals. 
The method described here (Sect. 3.5) reveals the near-total content and is 
aiming at determining the anthropogenic contamination. 

3.2 

Volatile Hydrocarbons 

■ Introduction 

Objectives. The volatile organic compounds ( VOCs) in soils primarily orig- 
inate from petroleum products and solvents. The spectra of the VOCs de- 
pend on their source. The analysis of benzene, toluene, and ethylbenzene 
and xylenes (BTEX) is widely used as an indicator of contamination with 
light petroleum products, e.g., petrol and kerosene. Furthermore, the gaso- 
line additives MTBE and TAME as well as halogenated volatile compounds 
can be analyzed with this method. 

Principle. A soil sample is extracted with methanol. A defined volume of the 
methanol extract is transferred into water and the water sample is heated 
to 80 °C in a headspace vial. When equilibrium is established between the 
gaseous and liquid phases, an aliquot of the gaseous phase is injected on 



100 K.S. j0rgensen et al. 

a column of a gas chromatograph and the VOCs are determined with a mass 
selective detector. 

Theory. VOCs are a group of compounds that have a boiling point from 20 
to 220 °C and usually they have two to ten C atoms. They are mainly un- 
substituted or substituted monoaromatics and short-chain aliphatic com- 
pounds that differ in solubility and in toxicity. The individual compounds 
are quantitatively determined using this method, as can also be the diaro- 
matic compound naphthalene. We do not recommend measuring the sum 
of VOCs because such a sum is unspecific and depends on the compounds 
included. 

The sampling (ISO 10381-1 1994; ISO 10381-2 1994; Owen and Whittle 
2003) is a crucial step in the analysis of VOCs. In order to prevent their loss 
during preparative steps, field-moist samples are used (ISO 14507 2003). 
The sample is added into a pre-weighed glass container containing a known 
amount of methanol. To control the quality of the determination, field du- 
plicates, a procedural blank, and a control sample are analyzed. The two 
main methods of analysis of VOCs are static headspace/gas chromatogra- 
phy (e.g., ISO/PRF 22155 in prep.) and purge and trap/gas chromatography 
(e.g., ISO 15009 2002). In the analysis of volatile aliphatic and aromatic 
hydrocarbons a mass selective detector (MSD) is used. VOCs can also be 
detected with a photo ionization detector (PID), a flame ionization detec- 
tor (FID), and an electron capture detector (ECD; Owen and Whittle 2003). 
The identification of target compounds (ISO/DIS 22892 in prep.) is easy 
with a MSD, and a possible matrix effect can be eliminated. The method 
described here is that using static headspace/gas chromatography (MSD) 
and is based on the proof of a new international standard ISO/PRF 22155 
and has earlier been described by Salminen et al. (2004). 

■ Equipment 

• Usual laboratory glassware, free of interfering compounds 

• Shaking machine 

• Headspace analyzer and gas chromatograph with a mass selective detec- 
tor (MSD) 

- Oven temperature program: maintain 35 °C for 2min, then steadily 
raise by 14°C/min up to 90 °C. Maintain 90 °C for 5min, then raise 
by 12°C/min up to 190 °C. Maintain 190 °C for 1 min, then raise by 
40 °C/min up to 225 °C, and maintain at 225 °C for 1 min. 

- Carrier gas: helium. 

- Gas flow: lOmL/min. 

- Split ratio (gas flow rate through split exit: column flow rate): 5.7:1. 



3 Quantification of Soil Contamination 101 

• Column: stationary phase non-polar or low polar fused silica capillary 
column; film thickness 1.4 p.m; column length 30 m; internal diameter 
0.25 mm 

■ Reagents 

• Methanol 

• Internal standards, e.g., toluene-d 8 , a,a,a-trifluorotoluene 

• Helium 

• Synthetic air 

• Volatile aromatic and halogenated hydrocarbons for standard solutions: 
MTBE, TAME, benzene, ethylbenzene, toluene, m-xylene, p-xylene, 
o-xylene, styrene, naphthalene, dichloromethane, chloroform, carbon 
tetrachloride, 1 ,2-dichloroethane, 1,1,1 -trichloroethane, cis- 1 ,2-di- 
chloroethene, trichloroethene, tetrachloroethene, chlorobenzene, 1,2- 
dichlorobenzene, 1 ,4-dichlorobenzene, 1 ,2,3 -trichlorobenzene, 1 ,2,4- 
trichlorobenzene, 1 ,3,5-trichlorobenzene 

• Standard stock solutions 

- Standard solutions: for each analyte, lOmg/mL of methanol 

- Internal standard (see above) solution, lOmg/mL of methanol 

• Working standard solutions 

- Standard solutions: 1 mg mixed analyte solution/mL of methanol 

- Internal standard (see above) solution, 10p.g/mL of methanol 

• Calibration solutions: at least five different concentrations by suitable 

dilutions of the working standard solutions within the range of 0.05- 
lOpg/L 

■ Sample Preparation 

In the field, approximately 20 g of field-moist soil sample is taken directly 
into a pre-weighed headspace vial containing 20 mL of methanol. No sieving 
of the samples is recommended. A separate sample is taken for dry mass 
determination in a glass jar leaving no headspace. 

■ Procedure 

1. Weigh the vial containing the soil sample and methanol. 

2. Shake the vial containing sample and methanol for 30 min with the 
shaking machine. 

3. Allow the vial to stand for 10-15 min to settle the solid material. 



102 K.S. j0rgensen et al. 

4. Pipette 10 mL of water, 100 pi of methanol extract, and 5pL of the 
working internal standard solution into a headspace vial. 

5. Place the vial in the headspace system and heat the sample at 80 °C for 
lh. 

6. Use headspace injection for gas chromatographic analysis. 

7. Detect the compounds with the mass selective detector (MSD). 

8. Identify the peaks of the internal standards by using the absolute re- 
tention times. 

9. Determine the relative retention times for all the other relevant peaks 
in the gas chromatogram. These retention times should be determined 
in relation to those of the internal standards. 

10. Determine the dry mass content, e.g., by using the method described 
in ISO 11465(Chapt.2) 

11. Calculate the concentrations of the analytes. 

To prepare a calibration curve, treat the calibration standards as the soil 
samples: 

1. Add 100 pL of calibration solution to a headspace vial containing 10 mL 
of water. 

2. Add a known amount of working internal standard solution into the vial. 

3. Close the vial and treat it according to the procedure. 

■ Quality Control 

1. Procedural blank determination: add 100 pL of methanol and 5 pi of the 
working internal standard solution to 10 mL of water. Treat this mixture 
as the soil sample. 

2. Control sample determination: add a known amount of working stan- 
dard solution to a pristine soil sample that contains neither VOCs nor 
methanol. Treat the control sample as the soil sample and calculate the 
recovery (%) of the analytes. Mark the recovery on the quality- control 
chart. 

■ Calculation 

Concentration of analytes is quantified with respect to the internal standard 
using the following formula: 

_ C iw X Vte X V, 
Cm,i — TA 

m dm x V, 



(3.1) 



a 



3 Quantification of Soil Contamination 103 

c m> i content of the analyte cc i" in the sample (mg/kg soil dry mass) 

C[ w mass concentration of the analyte "i" in the spiked water sample 
obtained from the calibration curve (pg/L) 

Vte total volume of the extract (methanol added to the soil sample + water 
in the sample obtained from the determination of dry mass content; 

mL) 

V w volume of the spiked water sample for headspace measurement (mL) 

^dm dry mass of the test sample used for extraction (g) 

V a volume of the aliquot of methanol extract used for the spiking of 
water sample for headspace measurement (pi) 

■ Notes and Points to Watch 

• Assure that compounds do not evaporate during sample handling. 

• Exposure of samples to air, even during sampling, shall be avoided as far 
as possible. 

• The use of plastics, other than PTFE, shall be avoided. 

• Samples shall be analyzed as soon as possible. 

• Store the samples in the dark at 4 ± 2 °C no longer than 4 days. 

• The standard and calibration solutions can be stored for 1 year at -18 °C. 

• The internal standard solutions can be stored for several years at -18 °C. 

• Avoid direct skin contact and inhalation of vapors from standards and 
samples. 

3.3 

Hydrocarbons in the Range Ci to C 40 

■ Introduction 

Objectives. Petroleum derivatives such as diesel fuel, heating oil, and lu- 
brication oil are widely used in human activities and thus are common 
pollutants in the soil environment. These petroleum products are complex 
mixtures of hundreds of various hydrocarbons. The analytical method 
described here (modified ISO 16703 2004 Salminen et al. 2004) allows 
a quantitative and a composition pattern determination of all hydrocar- 



104 K.S. j0rgensen et al. 

bons (that is, n-alkanes from CnH 2 2 to C 39 H 80 , isoalkanes, cycloalkanes, 
alkyl benzenes, and alkyl naphthalenes) with a boiling range of 196 to 
518 °C. Gasolines cannot be quantified using this method. Furthermore, 
high concentrations of polyaromatic hydrocarbons (PAHs) may interfere 
with the analysis. 

Principle. A soil sample is extracted by sonication with n-heptane-acetone 
including the internal standards (n-decane and n-tetracontane). To sepa- 
rate the organic phase, water is subsequently added. The extract is washed 
with water and the polar constituents and water are removed from the ex- 
tract with Florisil (U.S. Silica Co., Berkeley Springs WV, USA) and sodium 
sulfate, respectively. Hydrocarbons in the range from C 10 to C 40 are deter- 
mined from an aliquot of the purified extract with a gas chromatograph 
equipped with a flame ionization detector (FID). For the quantification of 
all the hydrocarbons in this range, the total peak area between the internal 
standards n-decane and n-tetracontane is measured. 

Theory. Petroleum derivatives are complex mixtures of various hydrocar- 
bons with different characteristics (e.g., volatility, water solubility, biode- 
gradability). In the assessment of petroleum hydrocarbon contamination 
and the effects of microbial activity (past, present, or future) on the fate 
of these contaminants in soil, it is essential to know the quantity and the 
composition of the contaminating agents. This information is of high value 
when, for instance, a bioremediation process is followed over a span of 
time. Moreover, as hydrocarbons differ in their amenability to microbial 
degradation, this information is of a remarkable value. 

In the past, gravimetric or infrared spectrometric methods have been 
extensively used for the determination of hydrocarbons in soil. While these 
methods can be used for quantification of a range of hydrocarbons, they 
do not provide any information of the their quality, that is, of their com- 
pound composition pattern. To obtain this information, more sophisticated 
methods such as gas chromatographic analyses, are employed. 

The extraction of hydrocarbons shall be performed in such a manner 
that the broad spectrum of the compounds of interest is included in the 
analysis. Moreover, it is essential that the extraction procedure is suitable 
for field-moist soil samples in which hydrocarbons may be attached to 
soil particles, and in which soil water present in the samples may impede 
the extraction of the non-polar hydrocarbons. Thus, a mixture of polar 
(acetone) and non-polar (n-heptane) solvents is used. On the other hand, 
polar compounds have to be removed from the extract as they interfere 
with the gas chromatographic analysis, and to avoid the inclusion of po- 
lar compounds other than petroleum hydrocarbons in the analysis. It is 
to be noted that PAHs and volatile compounds have to be analyzed sepa- 
rately. 



3 Quantification of Soil Contamination 105 

■ Equipment 

• Usual laboratory glassware free of interfering compounds 

• Sonicator 

• Laboratory centrifuge 

• Gas chromatograph (GC) with a non-discriminating injection system 
and a flame ionization detector (FID), helium as a carrier gas 

• Pre-column (in case on-column injection is used) 

• Capillary column specifications: 5% phenyl polysilphenylene-siloxane 
stationary phase, e.g., SGE BPX5 capillary column, 5 m length and 1.4- 
l^m film thickness 

■ Reagents 

• n-Heptane 

• Acetone 

• Ion- exchanged water 

• n-Decane (Ci H 22 ), n-eicosane (C 2 oH 42 ), n-triacontane (C 30 H 62 ), n-pen- 
tatriacontane (C 35 H 72 ), and n-tetracontane (C 40 H 82 ) - n-decane and n- 
tetracontane being used as the integration window and the latter also as 
an internal standard 

• Florisil (150-250 p.m, 60-100 mesh) (Activated Florisil is stored in a des- 
iccator and is usable for a week after the activation. Note: the activity of 
Florisil will gradually decrease after the activation.) 

• Anhydrous sodium sulfate (Na 2 S0 4 ) must be kept at 550 °C for at least 
2 h prior to its use 

• Diesel fuel and lubrication oil standards free of additives 

• Helium 

• Hydrogen 

• Synthetic air 

• Control soil sample 

• Standard stock solutions 

- Standard extraction solution (0.15 mg/mL of C 10 H 22 and 0.20 mg/mL 
of C 40 H 82 ): weigh 20 p.L of n-decane and 20 mg of n-tetracontane and 
dissolve in 100 mL of n-heptane. Prepare the solution in a volumetric 



106 K.S. j0rgensen et al. 

flask by weighing and calculate the accurate concentrations of the 
internal standards n-decane and n-tetracontane in the solution. Store 
the solution at 4 °C in the dark. The solution is usable for at least 6 
months if stored in a tightly closed (Teflon-capped) glass vial. 

- Working standard extraction solution: dilute the standard extraction 
solution 1:9 (v/v) in n-heptane. Prepare the solution in a volumetric 
flask by weighing and calculate the accurate concentrations of the 
internal standards n-decane and n-tetracontane in the solution. The 
solution is usable for 1 week if stored in a tightly closed (Teflon- 
capped) glass vial. 

- Calibration stock solution (20 mg hydrocarbons/mL): Weigh 100 mg 
of diesel fuel and 100 mg of lubrication oil and dissolve in lOmL of 
n-heptane. Prepare the solution in a volumetric flask by weighing and 
store the solution at 4 °C in the dark. The solution is usable for at least 
6 months if stored in a tightly closed (Teflon-capped) glass vial. 

- Working calibration solutions: prepare at least five solutions with 
final hydrocarbon concentration ranging from 0.1 to 2-3 mg/mL. 
Prepare the solution by diluting the calibration stock solution with 
n-heptane to obtain a final volume of 10 mL. Weigh the amounts of 
solutions used to calculate the exact hydrocarbon concentrations in 
the working calibration solutions. The solution is usable for at least 
6 months if stored in a tightly closed (Teflon-capped) glass vial. 

- Stock solution for testing the performance of the gas chromatograph: 
weigh 5.0 mg each of n-decane (C10H22), n-eicosane (C 2 oH 42 ), n-tria- 
contane (C 30 H 62 ), n-pentatriacontane (C35H72), and n-tetracontane 
(C 40 H 82 ) and dissolve them in 10 mL of heptane. Prepare the solution 
in a volumetric flask by weighing the mass of the added heptane to 
calculate the exact concentration of the individual n-alkanes in the 
solution. Store the solution at 4 °C in the dark. The solution is usable 
for at least 6 months if stored in a tightly closed (Teflon capped) glass 
vial. 

- Working solution for testing the performance of the gas chromato- 
graph: dilute the test stock solution in n-heptane in a ratio of 1:9 (v/v). 
Prepare the solution in a volumetric flask by weighing to calculate the 
exact concentration of the individual n-alkanes in the solution. 



■ Sample Preparation 

Sampling should be performed according to good practices (ISO 10381-1 
1994; ISO 10381-2 1994). For the analysis, a homogenized field-moist soil 



3 Quantification of Soil Contamination 107 

sample is used (ISO 14507 2003). However, if the water content of the sample 
is extraordinarily high, separation of the organic phase may occur prior to 
the extraction (that is, at the time of the introduction of the sample into the 
extraction solution). In such case, the sample has to be pre-dried overnight 
at room temperature prior to the extraction. 

■ Procedure 

Prior to Analysis 

1. Calibrate the gas chromatograph by running aliquots of the working 
standard solutions. 

2. An aliquot of the working test solution should be run on the GC and the 
yields of the individual n-alkanes calculated. The ratio between C 2 oH 42 
and C 40 H 82 should not exceed 1.2. 

Analytical Procedure 

1. Weigh 10 g of a sample into an extraction vial. 

2. Weigh 5-10 g of a control sample with a known concentration into 
a separate vial. 

3. Add 10 mL of working standard solution and 20 mL of acetone into each 
of these vials. 

4. Prepare a blank determination: add 1 mL of working standard solution 
and 20 mL of acetone but omit the sample. The blank and the control 
sample are treated in a similar manner to the (unknown) samples. 

5. Mix the samples gently and sonicate for 30 min. Add ice into the soni- 
cator to keep the samples cool. 

6. Add 30 mL of water and shake for 1 min. 

7. Centrifuge the samples (2,500 rpm, 5 min). 

8. Transfer the organic phase into a 25-mL test tube with a Teflon-lined 
screw cap, add 10 mL of water, and shake for 1 min. 

9. Transfer the organic phase into another test tube with a Teflon-lined 
screw cap and add approx. 0.5 g of Na 2 S0 4 and shake. 

10. Add approx. 1.5 g of Florisil into the tube and shake for 10 min in 
a mechanical shaker. 

11. Centrifuge the tubes (2,000 rpm, 1 min). 

12. Transfer an aliquot of the purified extract into a GC vial. Avoid the 
introduction of Florisil into the GC vial. 



108 K.S. j0rgensen et al. 

13. Run all the samples by GC. 

14. Solvent blank should be subtracted from the sample chromatogram. 
Integrate the total area between the peaks of Ci H 2 2 and C 40 H 82 to obtain 
the hydrocarbon concentration of the extracts from the calibration 
extracts. 

15. Integrate the total area of the C 40 H 82 peak to obtain the recovery of 
C 40 H 82 in the analysis. 

■ Quality Assurance 

1. The hydrocarbon concentration in the blank extract should be below 
0.025 mg/mL. 

2. The recovery of the internal standard n-tetracontane should be calculated 
in each extract. The yield should be 100 ± 20% of the theoretical value 
of C 40 H 82 in the extraction solution. 

3. The hydrocarbon content of the control soil sample should be monitored 
over time and the results ought to be analyzed according to general good 
quality procedures. 

■ Calculation 

The concentration of hydrocarbons in the range from C 10 H 2 2 to C 40 H 82 
(c H c) i n the sample is calculated as follows: 

c 2C x 10 x 1000 x / 

Chc = -£ - A (3.2) 

m x d s 

chc concentration of hydrocarbons in the range from Ci H 22 to C 40 H 82 
in the sample (mg/kg dry mass) 

c gc hydrocarbon concentration of the extract calculated from the cali- 
bration equation (mg/mL) 

10 volume of the organic solvent used in the extraction (10 mL of hep- 
tane) 

1,000 conversion factor of the soil mass (1 kg = 1,000 g) 

/ dilution factor (if applicable) 

m wet mass of the sample (g) 

d s content of dry substance in the field-moist sample (g/g), determined 
according to ISO 11465 (1993) 



3 Quantification of Soil Contamination 109 

■ Notes and Points to Watch 

• The samples should be analyzed as soon as possible. If this is not feasible, 
the samples should be stored at -20 °C. 

• Hydrocarbons are subjected to biodegradation both under aerobic and 
anaerobic conditions. Therefore, storage of the samples at temperatures 
above °C should be avoided (Salminen et al. 2004). 

• The efficacy of each Florisil stock has to be tested prior to its use in the 
analysis. 

• Weighing of the liquid, viscous standard compounds gives very precise 
solutions. 

• Avoid skin contact and inhalation of vapors from standards and samples. 

3.4 

Polyaromatic Hydrocarbons (PAHs) 

■ Introduction 

Objectives. Polycyclic aromatic hydrocarbons (PAHs) are often found at 
contaminated sites, particularly in connection with tar contamination at 
former gasworks. They also exist as diffuse contamination in urban areas 
and alongside roads. Furthermore, wood impregnation with creosote and 
incomplete combustion of hydrocarbons are major sources of PAHs in soil. 
Polyaromatic hydrocarbons are a group of more than 100 different com- 
pounds. This method describes the determination of a small selection of 
the many PAHs found in the environment. The US Environmental Protec- 
tion Agency (EPA) has chosen 16 of these PAHs to be the most important 
ones to be analyzed (EPA priority list 1982). 

Principle. A field-moist sample is extracted twice with acetone, and then 
hexane is added to the acetone extract. The extract is washed twice with 
water and the organic layer is dried with anhydrous sodium sulfate. When 
necessary, the extract is cleaned up by adsorption chromatography on 
a silica gel. The (purified) extract is analyzed by capillary gas chromatog- 
raphy with mass selective detection, using appropriate deuterated PAHs as 
internal standards. 

Theory. Polycyclic aromatic hydrocarbons occur ubiquitously in the en- 
vironment. Sixteen PAHs (Table 3.1) were chosen by the US EPA to be 
analyzed in environmental samples because they are the most abundant at 
hazardous waste sites and more information is available on these than on 
other PAHs. Moreover, the chosen compounds exhibit harmful effects that 



110 



K.S. j0rgensen et al. 



Table 3.1. Native and deuterated PAHs with their specific ions (target ion with qualifier ion 
in parentheses) 



Native PAH 


Mass number 


Deuterated PAH 


Mass number 




(amu) 




(amu) 


Naphthalene 


128(129) 


D 8 -Naphthalene 


136 


Acenaphthene 

Acenaphthylene 

Fluorene 


154(153) 
152(151) 
166(165) 


Dio -Acenaphthene 


164 


Anthracene 
Phenanthrene 
Fluoranthene 
Pyrene 


178 (89) 
178(179) 
202(101) 
202(101) 


D io -phenanthrene 


188 


Benz(a)anthracene 
Chrysene 


228(114) 
228(114) 


Di2-chrysene 


240 


Benzo(b)fluoranthene 
Benzo(k)fluoranthene 
Benzo(a)pyrene 
Indeno(l,2,3-cd) 


252 (253) 
252 (253) 
252 (253) 
276(138) 


D 12 -perylene 


264 


pyrene 

Dibenzo(ah)anthracene 

Benzo(ghi)perylene 


278(139) 
276(138) 







are representative of PAHs and exposure to these is more frequent than that 
to other PAHs. 

The most common analytical methods are based on liquid chromatogra- 
phy with fluorescence detection or UV detection (HPLC/FL or UV), or on 
gas chromatography with mass selective detection (GC/MSD). Both tech- 
niques have their benefits. Generally, HPLC has less resolution, which is 
problematic when the studied samples contain complex PAH mixtures. On 
the other hand, the UV and the fluorescence detection are highly sensitive 
and specific. Mass spectrometry is a powerful tool for identifying individ- 
ual compounds. The sensitivity of the GC/MSD can be increased if the mass 
spectrometer is operated in selected ion monitoring (SIM) mode. In the lit- 
erature, there are numerous applications available for analyzing PAHs that 
differ as to, e.g., extraction techniques, solvents used, and clean-up meth- 
ods. The method presented here is based on GC/MSD technology and on 
the draft international standard method (ISO/DIS 18287 in prep.), but uses 
hexane instead of petroleum ether as the solvent. The method is relatively 
fast and is applicable to all types of soils, covering a wide range of PAH 
contamination. With this method, other PAHs than those in the Table 3.1 



3 Quantification of Soil Contamination 111 

can be determined as well. A detection limit of 0.01 mg/kg dry mass can be 
ensured for each PAH. 

■ Equipment 

• Usual laboratory glassware free of interfering compounds 

• Shaking machine 

• Laboratory centrifuge 

• Gas chromatograph (GC) with a mass selective detector (MSD) 

- Oven temperature program: maintain 60 °C for 2min, then steadily 
raise by 20 °C/min up to 180 °C, then raise by 8 °C/min up to 280 °C 
and keep at that temperature for 10 min. 

- Splitless injection (split closed for 2 min) of 1 p.L. 

- Carrier gas: helium 1 mL/min. 

• Capillary column specifications: medium polar stationary phase, e.g., 
HP-5MS, film thickness 0.25 p.m, length 30 m, internal diameter 0.25 mm. 

■ Reagents 

• Acetone 

• n-Hexane 

• Ion- exchanged water 

• Anhydrous sodium sulfate (Na 2 S0 4 ), must be kept at 550 °C for at least 
2 h prior to its use 

• Silica gel 60 (particle size 60-200 pm), deactivated (Heat silica gel 60 for 
5h at 130 °C in a drying oven. Allow to cool down in a desiccator and 
add 10% water (w/w) in a flask. Shake for 5 min by hand until all lumps 
have disappeared, and then shake for 2 h in a shaking machine. Store 
deactivated silica gel in absence of air. It can be used for 1 week.) 

• Helium 

• Nitrogen 

• Quality control soil sample (e.g., certified reference material or in-house 
reference material) 

• Calibration stock solutions 

- Native PAHs (PAHs to be determined): commercially available cer- 
tified standard stock solution can be used with a concentration of 
approx. 100p.g/mL for each native PAH (e.g., Dr. Ehrenstorfer PAH 
Mix 9, X20950900CY). 



112 K.S. j0rgensen et al. 

- Deuterated PAHs (internal standards): commercially available cer- 
tified standard stock solution can be used with a concentration of 
approx. 1,000 pg/mL for each deuterated PAH (e.g., Dr. Ehrenstorfer 
PAH Mix 31, YA20953100TO). It is recommended that at least five 
deuterated PAHs be used as internal standards. The internal stan- 
dards are chosen to resemble the physical and chemical properties of 
the compounds to be analyzed (see Table 3.1). 

• Calibration working solution 

- Native PAHs: transfer 5 mL of the calibration stock solution contain- 
ing the native PAHs stock solution into a 25-mL volumetric flask and 
fill up to the mark with hexane (20 pg/mL) 

- Deuterated PAHs: transfer 1 mL of the calibration stock solution con- 
taining the deuterated PAHs stock solution to a 25-mL volumetric 
flask and fill up to the mark with hexane (40 pg/mL) 

• Calibration standard solutions: prepare a series of calibration standards 
over a suitable range (e.g., 0.2-10 pg/mL) by transferring 0.1-5 mL of 
the native PAH calibration working solution into a 10-mL volumetric 
flask and fill up to the mark with hexane. Transfer 1 mL of the standard 
solution into a GC vial and add 100 pL of the deuterated PAH calibration 
working solution. Each of the calibration standards nominally contains 
4 pg/mL of each of the deuterated PAHs. However, laboratories should 
determine their own concentration range depending on the samples to 
be analyzed. 

■ Sample Preparation 

Sampling should be performed according to good practices (ISO 10381-1 
1994, ISO 10381-2 1994). For the analysis, a homogenized field-moist soil 
sample is used (ISO 14507 2003). Stones and other bigger materials ob- 
viously not contaminated should not be analyzed. Large particles with 
expected contamination should be reduced in size and analyzed with the 
finer sample material. 

■ Procedure 

Extraction Procedure 

1. Weigh 10 g of a field-moist (or air-dried-overnight) sample into an ex- 
traction flask equipped with a Teflon inlay (a conical flask or a centrifuge 
tube with a capacity of 100 mL). 

2. Add 25 mL of acetone. 



3 Quantification of Soil Contamination 113 

3. Close the flask with a screw cap and extract by shaking for 15 min in 
a shaking machine. 

4. After settling, separate the organic phase into a shaking funnel of 
500 mL either by decanting or by using a centrifuge (2,500 rpm, 5 min). 

5. Repeat the extraction with 25 mL of acetone. 

6. Add 50 mL of hexane to the combined acetone extracts, and remove the 
acetone and other polar compounds by shaking with 100 mL of water. 
Discard the water and perform another wash in the same manner. 

7. If necessary, concentrate the extract on a water bath at 40 °C to about 
10 mL using a gentle stream of nitrogen at room temperature. Record 
the final volume of the extract and dry the concentrated extract over 
anhydrous sodium sulfate. 

8. Transfer 1 mL of the dried extract into a GC vial and add 100 p.L of 
the deuterated PAH calibration working solution. The sample then 
nominally contains 4p.g/mL of each of the deuterated PAHs. 

9. Prepare a blank determination in a similar manner but without any soil 
sample. 

10. Perform an extraction of the quality control soil sample in the same 
manner as of the test sample. 

Clean-Up Procedure 

1. If necessary, the extract can be cleaned with a silica gel adsorption 
column. Prepare the column by placing a small plug of glass wool on the 
bottom of the column, add 4 g of deactivated silica gel and then about 
1 cm of anhydrous sodium sulfate to the top. 

2. Condition the column by eluting 10 mL of hexane. When the eluant 
reaches the top of the column packing, transfer an aliquot (lmL) of 
the concentrated extract containing the internal standards to the top 
of the column. Elute with 50 mL of hexane and collect the extract in 
a point-shaped test tube. 

3. Concentrate the purified extract in a water bath at 40 °C to about 1 mL 
using a gentle stream of nitrogen at room temperature. 

4. Transfer the purified extract into a GC vial. 

Gas Chromatographic Analysis 

1. Set the gas chromatograph in such a manner that optimum separation of 
the PAHs is achieved. Special attention should be paid to benzo(b)fluor- 
anthene and benzo(k)fluoranthene separation. 



1 14 K.S. j0rgensen et al. 

2. Run the working standard solutions and all the samples by a GC with 
mass selective detection in the scan mode (mass range from 50 to 
300 amu). 

■ Quality Assurance 

1. The blank measurement of the total method should be carried out with 
each series of soil samples. The PAH concentration in the blank should 
be carefully studied, and if traces of contamination are found, the source 
of contamination should be investigated. 

2. The quality control sample should also be analyzed with each series of 
soil samples. The results should be monitored over time and the results 
treated statistically. 

■ Calculation 

For the quantitative analysis, a calibration curve of the ratio of the PAH 
determined to the internal standard peak area against the mass of PAH in 
the sample injected is constructed using the data handling system. Prepare 
these calibration curves for each native PAH using the specific ions (target 
ion as the quantitation ion and another ion as the qualifier ion), and the 
appropriate deuterated PAH as an internal standard (see Table 3.1). 

The amount of PAH in the GC vial (A PA h in p.g/mL) can be obtained from 
the calibration curve. Hence, the concentration of the native PAH in the 
soil sample can be calculated by the following equation: 

c n = x V x / (3.3) 

m x d s 

c n content of an individual PAH in the sample (mg/kg soil dry mass) 

Apah amount of PAH in the GC vial, obtained from the calibration curve 
(pig/mL) 

V volume of the concentrated extract (mL) 

/ dilution factor 

m mass of the sample (g wet mass) 

d s content of dry mass in the field-moist sample, determined according 
to ISO 1 1465 (g dry mass/g wet mass) 

■ Notes and Points to Watch 

• The samples should be analyzed as soon as possible. If not feasible, the 
samples should be stored at -20 °C. 



3 Quantification of Soil Contamination 115 

• Certain PAHs are carcinogenic and all the samples and standard solu- 
tions should be handled with extreme care. 

• The efficacy of each silica gel batch has to be tested prior to its use in the 
analysis. 

• For highly polluted soil samples, clean-up and concentration steps may 
not be necessary. 

3.5 

Heavy Metals 

■ Introduction 

Objectives. Most heavy metals are of geological origin, but contamination 
with them may be due to industrial, mining, agricultural, waste handling 
or other activity. Often a mixture of such metals occurs. The most common 
contaminants are arsenic, cadmium, chromium, copper, cobalt, lead, mer- 
cury, nickel, uranium, and zinc. In contrast to organic contaminants, heavy 
metals cannot be degraded by microbes or plants. Thus the bioremediation 
strategy is based on the movement of metals, e.g., from soil to plants as 
in phytoremediation, or on bioloeaching (see Chapt. 6). Some metals can 
undergo microbial oxidation-reduction or become methylated. Different 
ionic species of a heavy metal may have different toxicity, e.g., As 3+ is much 
more toxic than As 5+ . The method described here gives total concentra- 
tion of each metal, but does not give any information on the speciation. 
For that purpose separation of the ionic species may be achieved, e.g., 
by ion chromatography, followed by induced plasma mass spectrometry 
(ICP-MS). 

Principle. Soil samples are freeze dried, homogenized, sieved, digested in 
cone. HN0 3 in a microwave oven, and analyzed using ICP-MS. 

Theory. Traditional methods for heavy metals' extraction have been based 
on digestion in aqua regia (ISO 1 1466 1995) before determination by atomic 
absorption spectrometry (AAS), or more recently by ICP-MS. Destruction 
with hydrofluoric acid (ISO 14869-1 2001) is being used for some metal 
samples, e.g., in geological research. These extraction procedures give the 
highest yield of the metal content in a soil sample. However, these agents 
pose occupational health risks and alternative digestion using HNO3 has 
become common. The yield obtained using this method has been consid- 
ered sufficient in many countries for the determination of contamination 
with heavy metals (Karstensen et al. 1998). The method described here 
employs digestion with HNO3 and analysis by ICP-MS and has earlier been 
described by Salminen et al. (2004). 



116 K.S. j0rgensen et al. 

ICP-MS is a mult i- element analytical technique that can be used to 
measure the concentration of several elements simultaneously. The sample 
solution is nebulized into the plasma. A large percentage of atoms are 
ionized and a fraction of these ions are captured in the interface region of the 
system and channeled into the mass spectrometer. The mass spectrometer 
serves as a mass filter, and selectively transmits ions according their mass- 
to -charge ratio. 

The common elements to be analyzed by ICP-MS in soils are Al, As, Cd, 
Cr, Cu, Mn, Ni, Pb, Zn, B, Ba, Cs, Fe, Se, Sr, Ti, U, and V. Mercury is best 
determined by using the technique of direct combustion, which decom- 
poses the sample in an oxygen-rich environment and removes interfering 
elements. A dual-path-length cuvette/spectrophotometer specifically de- 
termines mercury over a wide dynamic range. The method for mercury 
requires no pretreatment other than freeze-drying, but a special piece of 
equipment is needed (e.g., an AMA254 Advanced Mercury analyzer); it is 
not described here in further detail. 

■ Equipment 

• Freeze drier 

• Microwave oven with Teflon tubes and a cooling system 

• Inductively coupled plasma mass spectrometer (ICP-MS) 

• Centrifuge and centrifuge tubes 

• Polystyrene tubes 

■ Reagents 

• Water: grade 1 as specified in ISO 3696 

• Digestion solution: cone. HN0 3 density 1.42 kg/L (69%) 

• Calibration standards: Single or multi-element (SPEX CertiPrep, Metu- 
chen, NJ, USA) 

- Multi-element solution 2(10 mg/L; Ag, Al, As, Ba, Be, Bi, Ca, Cd, Co, 
Cr, Cs, Cu, Fe, Ga, In, K, Li, Mg, Mn, Na, Ni, Pb, Rb, Se, Sr, Tl, U, V, 
Zn). Standards are commercially available in 5% HN0 3 . 

- Multi-element solution 4(10 mg/L; B, Ge, Mo, Nb, P, Re, S, Si, Ta, Ti, 
W,Zr) 

• Optimization solution: Mg, Ba, Rh, Pb, and Ce in 1% HN0 3 (10 }ig/L) 

• Internal standard: rhodium (1 mg/L) 



3 Quantification of Soil Contamination 117 

• Control material (NIST, Gaithersburg, MD, USA; SRM NIST 2709 San 
Joaquin Soil: Al, Ca, Fe, Mg, P, K, Si, Na, S, Ti, Sb, As, Ba, Cd, Cr, Co, Cu, 
Pb, Mn, Hg, Ni, Se, Ag, Sr, Th, V, Zn) 

■ Sample Preparation 

Use freeze-dried, homogenized, and sieved (< 2 mm) soil samples. 

■ Procedure 

1. Dry a frozen sample in a freeze-drier. 

2. Homogenize the dried sample manually. 

3. Sieve the sample (< 2 mm). 

4. Weigh accurately 0.25-0.5 g of the dried sample into a digestion tube. 

5. Add 5 mL of cone, nitric acid. 

6. Set one blank, one reference sample, and one duplicate sample to each 
batch. 

7. Digestion program: step 1: 250 W, 5min; step 2: 400 W, 5min; step 3: 
500 W, lOmin. 

8. Cool the digestion tubes to room temperature in a water bath. 

9. Open the tubes and transfer each digested solution quantitatively to 
a 30 mL plastic tube and dilute with water to a volume of 25 mL. 

10. If the samples are not clear, centrifuge at 3,000 rpm for 3 min. 

11. Dilute each sample (1:10 or 1:100) to a volume of 10 mL with water and 
add the internal standard (100 p.L of rhodium solution). 

12. The sample is ready for analysis. 

To perform a calibration, proceed as follows: 

1. Calibrate the instrument using two calibration solutions, namely, blank 
and 50p.g/L of standard solution. Normally multi-element standard so- 
lutions are used. 

2. Prepare 10 mL of the calibration solution and add the internal standard 
as described for soil samples. 

3. Perform the calibration and analyze the samples. 



118 K.S. j0rgensen et al. 

■ Calculation 

The mass concentration for each element is determined with the aid of the 
instrument's software. Enter the value of the dry mass of each sample into 
this software, and it calculates results directly in mg/kg soil dry mass. 

■ Notes and Points to Watch 

• Pay attention to the interference between/among different metals. 

• See that the acid concentration is same in the calibration and the sample 
solutions. 

• The instrument must be located in a laboratory free of contaminants. 

References 

Douglas DJ, Houk RS (1985) Inductively Coupled Plasma Mass Spectrometry. Prog Anal. 
Atom Spectrosc 8:1 

EPA Method 3015 (1994) Microwave assisted acid digestion of aqueous samples and ex- 
tracts for total metals analysis by FLAA, Furnace AA, ICP Spectrometry and ICP Mass 
Spectrometry 

ISO 10381-1 (1994) Soil quality - Sampling - Part 1: Guidance on the design of sampling 
programmes 

ISO 10381-2 (1994) Soil quality - Sampling - Part 2: Guidance on the design of sampling 
techniques 

ISO 11465 (1993) Soil quality - Determination of dry matter and water content on a mass 
basis - Gravimetric method 

ISO 11466 (1995) Soil quality - Extraction of trace elements soluble in aqua regia 

ISO 13877 (1998) Soil quality - Determination of polynuclear aromatic hydrocarbons 
(PAH) - Method using high performance liquid chromatography 

ISO 14507 (2003) Soil quality - Pretreatment of samples for determination of organic 
contaminants 

ISO 14869-1 (200 1 ) Soil quality - Dissolution for the determination of total element content - 
Part 1: Dissolution with hydrofluoric and perchloric acids 

ISO 15009 (2002) Soil quality - Gas chromatographic determination of the content of 
volatile aromatic hydrocarbons, naphthalene and volatile halogenated hydrocarbons - 
Purge-and-trap method with thermal desorption 

ISO 15587-2 (2002) Water quality - Digestion for the determination of selected elements in 
water - Part 2: Nitric acid digestion 

ISO 16703 (2004) Soil quality - Determination of content of hydrocarbon in the range Cio 
to C40 by gas chromatography 

ISO 17294-1 (2004) Water quality - Application of inductively coupled plasma mass spec- 
trometry (ICP-MS) - Part 1: General guidelines 

ISO 17294-2 (2003) Water quality - Application of inductively coupled plasma mass spec- 
trometry (ICP-MS) - Part 2: Determination of 62 elements 

ISO 3696 (1992) Water Quality - Water for analytical laboratory use - Specification and test 
methods 



3 Quantification of Soil Contamination 119 

ISO/DIS 18287 (in preparation) Soil quality - Determination of polycyclic aromatic hy- 
drocarbons (PAH) - Gas chromatographic method with mass spectrometric detection 
(GC-MS) 

ISO/PRF 22155 (in preparation) Soil quality - Gas chromatographic determination of the 
content of volatile aromatic and halogenated hydrocarbons and selected ethers - static 
headspace method 

ISO/DIS 22892 (in preparation) Soil quality - Guideline for GC/MS identification of target 
compounds 

ISO/TR 11046 (1994) Soil quality - Determination of mineral oil content - method by 
infrared spectrometry and gas chromatographic method 

j0rgensen KS, Puustinen J, Suortti A-M (2000) Bioremediation of petroleum hydrocarbon- 
contaminated soil by composting in biopiles. Environ Pollut 107:245-254 

Karstensen KH, Ringstad O, Rustad I, Kalevi K, j0rgensen K, Nylund K, Alsberg T, Olafs- 
dottir K, Heidenstam O, Solberg H (1998) Methods for chemical analysis of contaminated 
soil samples - tests of their reproducibility between Nordic laboratories. Talanta 46:423- 
437 

Laine MM, j0rgensen KS (1997) Effective and safe composting of chlorophenol- 
contaminated soil in pilot scale. Environ Sci Technol 31:371-378 

MLS- 1200 MEGA Microwave digestion system with MDR Technology; Operator Manual 
(1992) Milestone, Sorisole, Italy 

Owen S, Whittle P (2003) Volatile organic compounds. In: Thompson KC, Nathanail CP 
(eds) Chemical analysis of contaminated land. Blackwell Publ, CRC Press, pp 177-188 

Reference Manual Elan 5000 (1992) Perkin Elmer, Norwalk, Connecticut, USA 

Salminen JM, Tuomi PM, Suortti A-M, j0rgensen KS (2004) Potential for aerobic and anaer- 
obic biodegradation of petroleum hydrocarbons in boreal subsurface. Biodegradation 
15:29-39 

User's Manual Elan 5000 (1992) Perkin Elmer, Norwalk, Connecticut, USA 



4 



Immunotechniques as a Tool 
for Detection of Hydrocarbons 

Grazyna A. Plaza, Krzysztof Ulfig, Albert J. Tien 



4.1 

RaPID Assay Test System 

■ Introduction 

Objectives. Immunoassays (IMAs) are now being seen as useful analytical 
tools, and supplement to conventional analytical methods such as gas chro- 
matography and high performance liquid chromatography. The main IMA 
principle can be illustrated by the following reaction: Ab + Ag + Ag* ^> 
AbAg + AbAg* (Ab = antibody, Ag = antigen, Ag* = labeled antigen). 

Immunochemical methods provide rapid, sensitive, and cost-effective 
analyses for a variety of environmental contaminants (van Emon and 
Mumma 1990; van Emon and Lopez- Avila 1992; Marco et al. 1995). The 
driving force in the development of immunochemical methods is the need 
for rapid, simple, sensitive, and cheap tests that can be performed on- 
site without requiring sample transfer to an analytical laboratory. The 
increasing popularity of field IMA analyses can, in large part, be ascribed 
to portable equipment and minimal set-up requirements (van Emon and 
Gerlach 1995). Table 4.1 shows advantages and disadvantages of IMAs for 
environmental analyses. 

The following IMA techniques can be used in environmental studies: 
D TECH (Strategic Diagnostics, Newark, DE, USA), PETRORISC (EnSys, 
Research Triangle Park, NC, USA), EnviroGard (Millipore, Billerica, MA, 
USA), and RaPID (Ohmicron, Newtown, PA, USA) assays. Table 4.2 presents 
the properties of these systems and their application matrices and detection 
limits. 

Principle. The RaPID assay uses magnetic particles as the solid-support 
component of the ELISA (enzyme-linked immunosorbent assay). Attach- 
ing antibodies to microscopically small magnetic particles facilitates the 
chemical reaction between antibody and contaminant. The concentration 
of the compound to be detected is quantified after a color reaction. 

Grazyna A. Plaza, Krzysztof Ulfig: Institute for Ecology of Industrial Areas, 40-844 Katowice, 
6 Kossutha, Poland, E-mail: pla@ietu.katowice.pl 

Albert J. Tien: Holcim Group Support Ltd Corporate Social Responsibility Occupational 
Health and Safety, Im Schachen, 5113 Holderbank, Switzerland 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



122 



G.A. Plaza et al. 



Table 4.1. Some advantages and disadvantages of IMAs for environmental analysis (accord- 
ing to Sherry 1992; Sherry 1997) 



Advantages 



Disadvantages 



Wide applicability 

Sensitive, specific, and highly selective 

Rapid and easy to use 

Reduced preparation 

Rapid with high sample throughput 

Ideal for large sample loads; 
easily automated 

Suited to laboratory and field use 

Cost-effective analysis 
of small-volume samples 



Development costs 

Hapten synthesis can be difficult 

Can be vulnerable to cross reacting 
compounds and non-specific 
interferences 

Requires independent confirmation 

Not suited to small sample loads 
or multi-residue determinations 

Lack of acceptance 



Theory. One of the most common enzyme immunoassay (EIA) modifica- 
tions, sometimes termed "double antibody sandwich techniques," is ELISA 
(van Emon and Mumma 1990). ELISA is based on combining selective an- 
tibodies with sensitive enzyme reactions to produce analytical systems 
capable of detecting very low levels of chemicals. The RaPID system uti- 
lizes covalent binding of antibodies to magnetic particles that are made 
of silanized iron oxide (Fig. 4.1). The first stage is the immunochemical 
reaction between antibodies/magnetic particles and a chemical compound 
as antigen. The second stage is separation of magnetic particles from the 
antigen by applying a magnetic field. After washing, the color reagent is 
added and the concentration of the colored product is measured (RaPID 
assay environmental user's guide 1996; Plaza et al. 1999). The assay steps 
are presented in Fig. 4.2. 



Equipment 

RPA-I RaPID analyzer (spectrophotometer): laboratory bench-top- 
based, single wavelength, microprocessor-controlled analyzer 

Magnetic rack composed of two parts: the top rack holds the test tubes 
in place and the bottom base contains the magnets 

Portable balance 

Test tubes 

Vortex mixer 



Timer 



4 Immunotechniques as a Tool for Detection of Hydrocarbons 



123 



Table 4.2. A comparison of immunological test systems (EnviroGard Protocol 2004b, 
www.sdix.com) 







DTECH 


EnSys 


EnviroGard 


RaPID Assay 


Technology 


Latex 


Coated 


Coated 


Magnetic 






particle 


tube 


tube 


particle 


Result type 


Qualitative 


Qualitative 


Qualitative 


Qualitative 






and semi- 


and semi- 


and semi- 


and semi- 






quantitative 


quantitative 


quantitative 


quantitative 


Sample throughput 


1-4 


1-10 


1-17 


1-50 






samples/h 


samples/h 


samples/h 


samples/h 


Analy 


sis time 


20 min/run 


30 min/run 


30 min/run 


60 min/run 


EPA SW-846 method 


4030, 4035 


4030, 4035 


4030, 4035 


4030, 4035 


Storage 


' shelf life 


Ambient 


Ambient 


Refrigerated 


Refrigerated 






1 year 


1 year 


1 year 


1 year 


Training level 


Low; 


Medium; 


Medium; 


Medium; 






no training 


training 


training 


training 






required 


recommended 


recommended 


recommended 


Instrument 


DTECHTOR 


Photometer 


Photometer 


RPA-1 Analyzer 






Analyzer or 












color card 












Application matrix and detection limits 




Analyte 


Application 










BTEX 


Soil 


2.5-35 ppm 


— 


2 ppm 


0.9 ppm 




Water 


0.6-10 ppm 


— 


0.1 ppm 


0.09 ppm 


TPH 


Soil 


— 


10 ppm 


5 ppm 


10 ppm 




Water 


— 


— 


0.1 ppm 


1 ppm 


PAH 


Soil 


— 


1 ppm 


1 ppm 


0.2 ppm 




Water 


— 


15 ppb 


2 ppb 


0.9 ppb 


Carcino- 


Soil 


— 


— 


— 


10 ppb 


genic PAH 


Water 


— 


— 


— 


0.2 ppb 



Reagents 

All the reagents (Extraction Solution, Enzyme Conjugate, Antibody- 
Coupled Magnetic Particles, Color Reagent, Washing Solution, and Stop- 
ping Solution) are supplied by Ohmicron, Newtown, PA, USA, and their 
composition is under protection. 



Sampling and Sample Preparation 

Collect water and soil samples from the contaminated area in 500 mL 
wide-mouth bottles (Nalgene; Nalge Nunc, Naperville, IL, USA). 



124 



G.A. Plaza et al. 



TRADITIONAL 
IMMUNOASSAYS 



vs 



MAGNETIC PARTICLE-BASED 
TECHNOLOGY 



^7 

Antibody 




Antibodies are typically 
coated on plastic test tubes 
or plates 




^> 



Magnetic particle 
with antibody attached 



Antibodies are linking to 
microscopically small 
magnetic particles which 
speed up the chemical 
reaction between the antibody 
and the contaminant 



Fig. 4.1. Comparison of traditional IMAs and magnetic-particle-based technology (RaPID 
assay environmental user's guide 1996) 



Test directly samples or store them at 4 °C; water content in soil samples 
should not be more than 20-25%. 

Extract soil samples before testing: 

- Weigh 10 g of soil into the soil collection tube (Fig. 4.3) and add 20 mL 
of the extraction solution; screw the cap on tightly. 

- Shake vigorously and continuously for at least 60 s. 

- Remove the screw cap and attach the filter cap, then attach the plunger 
rod to the plunger of the soil collector, and filter the extract into the 
Extract Collection Vial. 

- Fill with 0.5-1 mL of the filtrate and cap the vial. 



■ Procedure 

1. Mix 200 pL of soil extract or water sample with 250 pL of the Enzyme 
Conjugate and 500 pL of antibody-coupled magnetic particles; incubate 
the mixture for 15 min at room temperature. 

2. Put all tubes into the magnetic rack and wait 2 min for the particles to 
separate. 



4 Immunotechniques as a Tool for Detection of Hydrocarbons 



125 



Immunological reaction 




o — «] — * 




Separation 



Stepl 




Color Development 




Step 3 



Legend 



o — < Magnetic particle with antibody 

<\ — * Antigen conjugate with 

^ Antigen 

□ Chromogen substrate 

■ Colored Product 



Fig. 4.2. Principle of RaPID assay (RaPID assay environmental user's guide 1996; Plaza et al. 
1999) 



Soil collector 




TOP _ Screw cap 



BOTTOM 



b 



T 




Luer cap 



Plunger rod Plunger 



Luer cone 



Fig. 4.3. Soil collection tube (RaPID assay environmental user's guide 1996) 



126 G.A. Plaza et al. 

3. Add 1 mL of washing solution, vortex each tube and wait 2 min. Repeat 
this step. 

4. Add 500 pL of color reagent. S^'^^-tetramethylbenzidine is used as 
the chromogen. Incubate for 20 min at room temperature. 

5. Add 500 pL of stopping solution (0.5% sulfuric acid). 

6. Within 15 min after adding the stopping solution, transfer 1 mL of solu- 
tion to cuvettes and read absorbance of standard solutions and samples 
at 450 nm using the RPA-I analyzer. 

■ Calculation 

1. The concentration of the colored product is directly proportional to the 
concentration of the labeled compounds. 

2. The RPA-I analyzer can perform mathematical computations, and sam- 
ple concentrations with statistics are obtained. Results are directly re- 
ported in ppb (ng/g) or ppm (pg/g). 

3. The hydrocarbon concentration in soil is calculated according to the 
following formula, taking into account the concentration calculated by 
the analyzer (ppb or ppm), the volume of extraction solution (20 mL), 
and the mass of soil used for extraction (g dry mass): 

concentration x volume 

Hydrocarbon concentration (ppb or ppm) = 

soil mass 

■ Notes and Points to Watch 

• Temperature control is required for reagent storage (4-8 °C) and during 
the performance of the assay (room temperature: 15-30 °C). 

• Use specific test kits for specific hydrocarbons; do not mix the reagents. 

• All the reaction steps should be done exactly according to the assay 
protocol. 

• Do not use test kit components after the expiration date. 

4.2 

EnviroGard Test System 

■ Introduction 

Objectives. Environmental IMAs have been developed and evaluated for 
analyses including major classes of pesticides, organic compounds such 



4 Immunotechniques as a Tool for Detection of Hydrocarbons 127 

as polychlorinated biphenyls (PCB), polyaromatic hydrocarbons (PAH), 
pentachlorophenols (PCP), benzene/toluene/ethylbenzene/xylene (BTEX), 
total petroleum hydrocarbons (TPH), dioxins and furans, microbial tox- 
ins, as well as inorganic compounds such as cadmium, lead, and mer- 
cury (Vanderlaan et al. 1988; van Emon and Mumma 1990; Sherry 1992; 
van Emon and Lopez- Avila 1992; Knopp 1995; van Emon and Gerlach 
1995; Gerlach et al. 1997). The EnviroGard test systems are quick and 
reliable with semi-quantitative results allowing screening at various lev- 
els of contamination (Table 4.2). They can be used during site remedia- 
tion to detect contaminants and monitor the cleanup and industrial pro- 
cesses. 

Principle. EnviroGard uses coated polystyrene test tubes as the solid sup- 
port component of the ELISA. The system is based on the use of polyclonal 
antibodies, immobilized on the test tube walls, that can bind specific con- 
taminants. 

Theory. The EnviroGard IMA kit uses 12 x 75 mm polystyrene test tubes 
coated with an antibody against the target contaminant (analyte). Coated 
polystyrene test tubes allow screening for various contaminations like PCB, 
PAH, TPH, BTEX, and PCP. When analytes are present in the sample, they 
compete with the specific Enzyme Conjugate (labeled analyte) for a limited 
number of binding sites on the antibodies (EnviroGard Protocol 2004a, 
2004b). According to the principles of competitive IMAs, the absorbance 
signal (or optical density) of the final reaction mixture is inversely pro- 
portional to the concentration of the contaminant (analyte) present in the 
test sample. After the immunological reaction, the unbound molecules are 
washed away and a chromogenic substance is added to the test tube. In 
the presence of bound specific enzyme conjugate, the clear substance is 
converted to a blue color. 



Equipment 

SDI Sample Extraction Kit contains devices to process 12 samples, i.e., 
12 each: extraction jars with screw caps, filter modules, ampule crack- 
ers, wooden spatulas, weigh canoes, disposable transfer pipettes, and 
ampules each containing 10 mL of 100% methanol 

20 antibody-coated test tubes (12 x 75 mm), 1 vial of negative control 
(methanol), calibrator vials, 1 vial of Hydrocarbons-Enzyme Conjugate, 
1 vial of Substrate Solution, 1 vial of Stop Solution, pipettes and tips 

Portable balance 

Spectrophotometer 



128 G.A. Plaza et al. 

• Test tube rack 

• Timer 

■ Reagents 

• Methanol (100%) 

• Stop solution: 1 N HC1 

■ Sampling and Sample Preparation 

• Collect soil without excess twigs, rocks, or pebbles in labeled 500 mL 
plastic containers. 

• Dry soil samples with a water content of 30% or more (by mass) before 
testing. 

• Store soil samples at 4 ° C in tightly sealed containers to avoid evaporative 
losses. 

• Use 10 mL of methanol to extract hydrocarbons from 10 g soil. When 
extracting clay samples add an additional 10 mL of methanol to the 
sample and shake vigorously for 1-2 min. 

• Filter the extract using Whatman #1 filter paper. A clay sample may soak 
up all the methanol, leaving little or no excess liquid to filter. 

■ Procedure 

1. Incubate all reagents at room temperature (at least 1 h) before use. 

2. Remove the antibody-coated test tubes (20 tubes/assay) from the foil 
pouch and label as negative control (NC), calibrators (CI, C2, etc.), and 
samples (SI, S2, etc.). 

3. Place the test tubes in the test tube rack, and add 100 pi of sample extract. 

4. Add 200 pi of specific enzyme conjugate to all test tubes. 

5. Gently shake the rack to mix for 10-15 s, and leave the tubes undisturbed 
for 10 min. 

6. Shake out the test tube contents vigorously into a sink, and wash the test 
tubes with double distilled water. Repeat this wash step three times. 

7. Add 500 pi of substrate solution to all test tubes, briefly shake, and then 
incubate for 5 min at room temperature. 

8. Add 500 pi of stop solution to all test tubes. 



4 Immunotechniques as a Tool for Detection of Hydrocarbons 129 

9. The results can be interpreted visually within 5 min after adding the 
substrate solution, or can be obtained more precisely with a spectropho- 
tometer. 



■ Calculation 

1. One enzyme molecule can convert many substrate molecules. Every 
test tube has the same number of antibody-binding sites and receives 
the same number of the specific enzyme conjugate molecules. Color 
development is inversely proportional to the hydrocarbon concentration, 
i.e., dark color indicates low concentration and light color indicates high 
concentration. 

2. Visual interpretation: compare the sample test tube to the calibrator test 
tubes against a white background. If a sample test tube contains a darker 
color than the calibrator test tube, the sample contains hydrocarbons at 
a concentration lower than the calibrator. If the sample test tube is less 
colored than the highest calibrator test tube, the soil extract should be 
diluted in methanol and the assay performed again. 

3. Photometric interpretation: place the negative control test tube into the 
spectrophotometer. Record the optical density (OD) at 450 nm for the 
negative control, and then measure the OD of calibrator #1. Repeat this 
step to determine the OD for each of the remaining calibrators and for 
samples. 

An example of data is given here: 
Tube OD 450 nm Interpretation 



NC (negative control) 


1.00 




CI (2ppm) 


0.87 




C2(10ppm) 


0.52 




C3 (50 ppm) 


0.25 




SI (sample 1) 


0.45 


> 10 ppm, < 50 ppm 


S2 (sample 2) 


0.72 


> 2 ppm, < 10 ppm 



Notes and Points to Watch 

The EnviroGard test is a screening test only. 

The test system can be used in the laboratory and in the field. 

Store all test kit components at 4-8 °C when not in use. 

Storage all reagents at ambient temperature (18-27 °C) on the day before 
using. 



130 G.A. Plaza et al. 

• Use only reagents or test tubes from one kit; do not mix the components 
from different test kits. 

• Do not expose substrate to direct sunlight. 

• Do not freeze test kit components or expose them to temperature greater 
than 37 °C. 

• Use gloves and protective clothing during the experiment. 



References 

EnviroGard™ Protocol (2004a) Petroleum fuels in soil. Test kit 70040000. Strategic Diag- 
nostics Inc, USA 

EnviroGard™ Protocol (2004b) Remediation assessment and industrial testing. Strategic 
Diagnostics Inc, USA (www.sdix.com) 

Gerlach RW, White RJ, O'Leary WFD, van Emon JM (1997) Field evaluation of an immunoas- 
say for benzene, toluene and xylene (BTX). Wat Res 31:941-945 

Knopp D (1995) Application of immunological methods for the determination of environ- 
mental pollutants in human biomonitoring. A review. Anal Chem Acta 311:383-392 

Marco MP, Gee S, Hammock BD (1995) Immunochemical techniques for environmental 
analysis. II Antibodies production and immunoassay development. Trends Anal Chem 
14:415-428 

Plaza G, Ulfig K, Bevolo AJ (1999) Application of the immunoassay techniques for the 
determination of PAHs and BTEX in soil. Int Environ Tech 9: 9-10 

RaPID assay environmental user's guide (1996), OHMICRON, USA 

Sherry J (1992) Environmental chemistry. The immunoassay option. Crit Rev Anal Chem 
23:217-300 

Sherry J (1997) Environmental immunoassays and other bioanalytical methods. Overview 
and update. Chemosphere 34:1011-1025 

Van Emon JM, Gerlach CL (1995) A status report on field-portable immunoassay. Environ 
SciTechnol 29:312-317 

Van Emon JM, Lopez- Avila V (1992) Immunochemical methods for environmental analysis. 
Anal Chem 64:79-88 

Van Emon JM, Mumma RQ (1990) Immunochemical methods for environmental analysis. 
ACS Symp Series 442, Am Chem Soc, Washington, DC 

Vanderlaan M, Watkins BE, Stanker L (1988) Environmental monitoring by immunoassay. 
Environ Sci Technol 11:247-254 



5 



Feasibility Studies for Microbial Remediation 
Hydrocarbon-Contaminated Soil 

Ajay Singh, Owen P. Ward, Ramesh C. Kuhad 



5.1 
Introduction 

While bioremediation processes are considered to be advantageous in terms 
of their relatively low cost, process flexibility, benign nature environmen- 
tally, and on-site utility, there have also been many instances where the 
processes have failed to achieve the required low contaminant concentra- 
tion criteria (Mandelbaum et al. 1995; Iwamoto and Nasu 2001; Grommen 
and Verstraete 2002). These failures have reduced consumer confidence 
in bioremediation and consequently the technology only garners a small 
portion of the US$ 7-8 billion US annual remediation market (Srinivasan 
2003). Bioremediation processes must comply with generally accepted good 
operating principles and have predictable end-points. The processes must 
be validated in advance such that they do not fail (Ward 2004). Feasibil- 
ity studies are therefore critical for the implementation of a successful 
bioremediation technology. 

Contaminated sites never exhibit identical characteristics and the expe- 
rience from one site can only be exploited at another to a limited extent. 
The biodegradation process in soil is complex, involving diffusion of con- 
taminants in the soil matrix, adsorption to the surface of soil particles, and 
biodegradation in the biofilms existing on the soil particles, in pores, and 
in the bound and free water after desorption from the soil surfaces. A va- 
riety of complex biodegradation patterns result from physical interactions 
between pollutants and soil matrix and from biological interactions among 
different organisms. Numerous factors such as soil moisture, pH, temper- 
ature, aeration, nutrient sources, type of soil, type(s) of contaminant(s), 
and interplay between these factors, affect the ecology of the microbial 
population and degradation of hydrocarbons in contaminated soil. 

For full-scale bioremediation applications, several important points need 
to be considered. In particular, how low can the concentration of the con- 



Ajay Singh, Owen P. Ward: Department of Biology, University of Waterloo, Waterloo, On- 
tario, Canada N2L 3G1, E-mail: ajasingh@sciborg.uwaterloo.ca 

Ramesh C. Kuhad: Department of Biotechnology, Kurukshetra University, Kurukshetra - 
136 119, Haryana, India 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



132 A. Singh et al. 

taminant be obtained during treatment considering: bioavailability and 
microbial activity, the fate of the contaminant in terms of mineralization, 
biotransformation, evaporation, build-up of microbial biomass, sorption 
to soil - and also considering indicators of time needed to obtain the set 
goal such as degradation rate for achieving the target level of contaminant, 
and finally the capital and operating costs? 

Both slurry bioreactors and land treatment technologies have success- 
fully been used to remove hydrocarbons from petroleum-contaminated 
soils. Naturally occurring or introduced microbial populations convert 
hydrocarbons to carbon dioxide, water, biomass, and humic material. For 
a successful bioremedial treatment of soil, it is important to consider several 
factors, including type and extent of contamination, bacterial population 
present, duration since contamination, optimal microbiological conditions, 
soil characteristics, proper bioremediation technique, and appropriate an- 
alytical method. Proper planning for the execution of a bioremediation 
technology based on the above criteria is of utmost importance to minimize 
risk of failure in terms of effort, time, and money. Protocols for conducting 
feasibility studies are, therefore, required to evaluate the effectiveness of 
a planned treatment method. 

In this Chapter, methods commonly used for determination of biodegra- 
dation potential in feasibility or biotreatability studies on bioremediation 
of contaminated soils and sludges are discussed. During feasibility stud- 
ies certain environmental and nutritional parameters are optimized for 
achieving accelerated and complete biodegradation of hydrocarbons. 

5.2 

Determination of Biodegradation Potential 

The data from the feasibility studies of hydrocarbon-contaminated soil is 
used to design a suitable full-scale bioremediation technology. Hence, it is 
important to carefully plan the biotreatability studies. Successive stages of 
planning and execution of feasibility studies are shown in Fig. 5.1. 

5.2.1 

Sampling and Soil Preparation 

Preparation of soil samples for site characterization or biodegradation 
experiments can alter the physico-chemical and biological properties of the 
original soil present in the field. In order to assess the extent of the overall 
soil contamination, proper field sampling procedures are required to obtain 
representative samples. The spatial variability of soil characteristics maybe 



5 Feasibility Studies for Microbial Remediation 



133 



Hydrocarbon-contaminated site/soil/sludge 



Assessment of contaminated waste or site and sampling 



Analysis of samples and characterization of waste 



Feasibility or biotreatability studies planning 



Microcosms or Mesocosms or Composting technique 



Determination of biodegradation potential 



Optimization of environmental and nutritional conditions 



Analysis and process monitoring 



Evaluation and interpretation of results 



Measurements of success 



Selection of the appropriate bioremediation technology 



Cost-benefit analysis 



Design and implementation of the technology 



Fig. 5.1. Successive stages for planning and implementation of feasibility studies 



considered for determination of the location and number of representative 
samples in heterogeneous and very large sites. The number of samples to be 
collected and analyzed usually depends on the overall sampling objective, 
contaminant distribution, size of the contaminated site and sampling and 
analytical costs (Huesemann 1994a). 

1. Samples may be collected from a uniform depth, using proper soil sam- 
pling equipment such as a core or a split-spoon sampler at 20-30 lo- 
cations and pooled. Top 10-15 cm of soil is often sampled. There are 
many procedures for choosing a sampling location. The most popular 
ones are either randomly selecting field locations for periodic sam- 
pling or placing a hypothetical square or rectangular grid over the site 
and taking samples at the center of each grid. Samples should be ho- 
mogenized prior to subsampling and submission for analysis (see also 
Chapt. 1). 

2. Soil samples are stored in either plastic bags or in glass or other non- 
reactive containers in a cooler on ice for immediate analysis. For long 
term storage, the samples should be at the field moisture levels and 



134 A. Singh etal. 

stored in glass or other non-reactive container around 4°C. Air drying 
of samples should be avoided. 

3. For biodegradation experiments, a relatively homogeneous sample 
should be prepared by sieving soil through a 2-4 mm sieve and soil 
moisture level determined so that all the experiments are carried out 
using the same conditions. Depending on the aim of the experiment, 
contaminant level, water content, pH, organic matter, nutrient levels, 
and type of soil are determined by standard methods (see Chapt. 2). 

5.2.2 

Selective Microbial Enrichment 

Successful application of a bioremediation technique requires the iden- 
tification of favorable conditions for the useful hydrocarbon-degrading 
microorganisms to actively grow and metabolize contaminants. Classical 
isolation methods, selective enrichment of specific microbes, and genetic 
approaches can be used to obtain a single microbial species or a group 
of different microorganisms (a consortium; see Chapt. 13). Selective en- 
richment is the most practical approach for large scale applications where 
the enrichment process is designed to increase the population of specific 
microorganisms. Suitable conditions and selective pressures are applied to 
encourage growth of microbes capable of degrading a particular substrate 
or a mixture of potential contaminants, in the growth medium, that are the 
targets for biodegradation or bioremediation as the sole sources of carbon 
(Vecchili etal. 1990). 

The initial inoculum can be obtained from the contaminated soil, sludge, 
or wastewater with known degradative activity. A consortium of hydrocar- 
bon-degrading microorganisms can be obtained by adding about 1 or 2 g 
of hydrocarbon-contaminated soil or sludge to 100 mL mineral medium in 
Erlenmeyer flasks. Culture can be further maintained in a flask by routinely 
transferring a 2% inoculum into a fresh medium at weekly intervals. 

Cyclone fermenters (Liu 1989) with a 1-L working volume can also be 
used to develop and maintain hydrocarbon-degrading cultures (Fig. 5.2). 
The cultures obtained by selective enrichment in the flasks can be inoc- 
ulated into a liter of mineral medium in a cyclone fermenter. The culture 
is maintained once weekly by removing about 50% of the volume of the 
culture and replacing with fresh medium and a known amount of the 
hydrocarbon as a sole carbon source. 

Soil columns may be used to enrich for hydrocarbon-degrading cultures. 
Glass soil columns with an inner diameter of 40 mm and length of 350 mm 
packed with 50 g air-dried soil and slightly moist pre-washed quartz sand 
between two layers of glass wool can be used to enrich desired strains 



5 Feasibility Studies for Microbial Remediation 



135 




Fig. 5.2. Cyclone fermenters used for main- 
tenance of hydrocarbon-degrading mi- 
croorganisms 



(Pfarl et al. 1990). The columns are rinsed with mineral medium several 
times prior to enrichment process and air is provided by using compressed 
air bubbled through distilled water at 1-2 L/h. 



5.2.3 
Controls 

In bioremediation experiments, sterile controls should frequently be used 
to demonstrate the biological activity and biodegradation process, since 
abiotic loss mechanisms such as adsorption and volatilization can occur 
simultaneously. Soil sterilization can be achieved by various physical and 
chemical methods. Methods include using chemicals such as mercuric 
chloride and sodium azide, as well by as autoclaving and providing gamma 
radiation. For inoculation of sterile soil, autoclave and gamma radiation 
are the most suitable because no residual chemical is left after sterilization. 
However, any sterilization method will alter the soil properties. Wolf et al. 
(1989) found that mercuric chloride had the least effect on soil properties 
such as pH, surface area, and release of Mn among various sterilization 
methods/agents tested and compared such as gamma radiation, microwave, 
dry heat, propylene oxide, sodium azide, mercuric chloride, chloroform, 
and antibiotics. 

In bioaugmentation studies, where the experiments are conducted with 
added inoculum to the soil, a killed-culture inoculum treatment should 
be included as control for possible nutrient effects of dead microbial cells. 
Mass balance of the existing or spiked contaminant should be determined 
at the conclusion of the experiment to evaluate contaminant disappearance 
due to biodegradation or abiotic mechanism. 



136 A. Singh et al. 

5.2.4 

Soil Microcosms 

One of the simplest methods requiring minimal equipment for soil biode- 
gradation studies is with use of a biometer flask (Bellco, Vineland, NJ, USA). 
The United Sates Environmental Protection Agency (US EPA) and Organi- 
zation of Economic Cooperation and Development (OCED) have also rec- 
ommended this method (OECD 1981; McFarland et al. 1991; Skladany and 
Baker 1994). Biodegradation activity can be evaluated by directly monitor- 
ing the loss of the target compounds or indirectly by measuring by-products 
of biodegradation or electron acceptor consumption. 

A biometer flask, a 250-mL Erlenmeyer flask with a side arm contain- 
ing potassium hydroxide to trap C0 2 evolved during biodegradation, is 
used in batch experiments to monitor degradation of the target compound 
present in or added to the contaminated soil. For biodegradation feasibil- 
ities studies, around 20% (w/v) aqueous soil suspension is recommended. 
Flasks are incubated with or without C0 2 -free air and periodically KOH 
solution is withdrawn and titrated with a standard acid solution to de- 
termine the amount of C0 2 produced. The matrix can be analyzed at the 
end of the test for organic and inorganic compounds. The biometer flasks 
can be modified to investigate specific problems related to specific types 
of contaminants and challenges in studying a given biodegradation. This 
flask system can be used to study biodegradation of both semi- volatile and 
volatile compounds, and to screen commercial inoculates as well. 

An electrolytic respirometer, designed to measure the oxygen uptake or 
rate of respiration by microbes in soil and sludge has been used by the 
US EPA for evaluation of commercial products for use in Prince William 
Sound, Alaska (Venosa et al. 1992). The respirometer consists of a reactor 
module connected to an electrolytic oxygen generator. The depletion of 
oxygen by microbes creates a vacuum that triggers the oxygen generator. 
The electricity used to generate the oxygen is proportional to the amount 
of oxygen (mg/L), while the C0 2 produced by microbial activity is trapped 
in KOH solution. The decision to choose a better amendment is based on 
high oxygen uptake rate, growth of degraders, and significant degradation 
of aliphatic and aromatic hydrocarbons. 

Another method to quickly determine biotreatability of hydrocarbon- 
contaminated soils and sludges is to simply use 250 mL Erlenmeyer flasks 
with working volumes of 50 mL containing 20% (w/v) soil or sludge slurry. 
For petroleum-contaminated soil or sludge samples, total petroleum hy- 
drocarbon (TPH) content is determined as hexane-extractable material. 

1. Set up at least 6 flasks for each test. 

2. Add a known mass of sludge or contaminated soil to the flask in order 



5 Feasibility Studies for Microbial Remediation 137 

to obtain less than 20% solids and 10% TPH concentration in a total 
working volume of 50 mL. 

3. Add 45 mL of the nutrient medium and 1.25 mL (0.25% final concentra- 
tion) of a non-ionic surfactant (10% w/v stock solution). 

4. Adjust pH of the contents to 6.8-7.2 using 5 N NaOH or 5 N HC1. 

5. Inoculate the flask with 2.5 mL (5%, v/v) of a microbial inoculum. 

6. Incubate the flasks at 30 °C for 14 days on a shaker (200 rpm). 

7. Extract the whole contents of 2 flasks with equal volume of n-hexane at 
the starting time of the test to determine initial TPH content. Extract 
contents of 2 flasks each with hexane after 7 and 14 days to determine 
residual TPH contents. 

8. After determination of TPH (Chapt. 3), dissolve the residue in a known 
volume of hexane for gas chromatographic analysis of hydrocarbons. 

In the above method, at least duplicate flasks should be set up for each 
sampling point and the contents of whole flasks should be extracted to 
determine residual hydrocarbons. Appropriate, controls for abiotic losses 
should be also be set up as described above. 

5.2.5 

Slurry Bioreactors 

The slurry bioreactor approach is to suspend and mechanically mix soil 
in aqueous solutions in a contained vessel or tank. Land-based systems 
usually require very long treatment times due to lack of control of envi- 
ronmental factors such as seasonal variation in temperature, pH, moisture, 
as well as of natural microbial activity, and mixing and circulation limita- 
tions. These problems can be eliminated in bioreactor systems, which are 
characterized by much higher rates and extents of degradation due to the 
minimization of mass-transfer, increased desorption of contaminants by 
continuous mixing, and control of environmental and nutritional factors 
such as pH, temperature, and moisture, bioavailability of nutrients and oxy- 
gen in order to promote rapid microbial growth and activity (Singh et al. 
2001; Van Hamme et al. 2003). 

Process conditions in bioreactors can be optimized for biodegradation 
depending on the nature of contaminant. Desired temperature and pH can 
be consistently maintained throughout the process and suitable amend- 
ments such as nutrients, surfactants, and microbial cultures can be sup- 
plied. Several examples of slurry reactors can be found in the literature. 



138 A. Singh etal. 

A method developed in the authors' laboratory and successfully scaled up 
for field applications is described here. 

1. Depending on the availability, use a 1-5 L or even larger volume biore- 
actor fitted with pH, temperature, and dissolved oxygen control for 
biotreatability studies. Alternatively, construct an inexpensive bioreac- 
tors by putting an air sparger in a glass or metal beaker or container. 

2. For biotreatability studies in the bioreactor, and depending on the soil 
and sludge composition, mix a sample of about 20% solids by mass 
with aqueous nutrient medium. 

3. Depending on the critical micellar concentration, add a non-ionic sur- 
factant with a hydrophilic-lipophilic balance (HLB) value 12-13 to ob- 
tain final concentration of 0.05-0.25%. 

4. Adjust the pH of the medium to around 7.0 using NaOH or HC1 solu- 
tions. 

5. Add the inoculum, prepared and maintained in a cyclone fermenter as 
described in Sect. 5.2.2, at the level of 10% (v/v) to the bioreactor. 

6. Maintain an aeration level of 0.1-0.2 wm (volume per volume per 
minute) during the process to avoid oxygen limitation in the system. 
Dissolved oxygen concentration should be maintained above 2 mg/L. 

7. A small mixer can also be used at about 200-300 rpm to achieve better 
mixing of the reactor contents. 

8. Keep the temperature at between 28 and 32 °C using a water bath or 
heater. 

9. Monitor the pH regularly and maintain it between 6.5 and 7.5 through- 
out the process. 

10. Compensate for any losses due to evaporation of water by adding water 
to the working volume level. 

11. Total microbial count and hydrocarbon-degrading bacteria can be de- 
termined at regular intervals to monitor the progress of biodegradation. 

12. Monitor biodegradation of hydrocarbons at periodic intervals for 2- 
4 weeks. 

The experimental design and data analysis during a biotreatability study 
will depend on the specific aim of the study. The slurry reactor experiments 
should be repeated to ensure consistent results. While sub-sampling over 
an extended period in a bioreactor experiment, care should be taken to 
ensure that the volume in the reactor is not drastically reduced. 



5 Feasibility Studies for Microbial Remediation 139 

5.2.6 

Land Treatment 

A set of laboratory experiments using contaminated soil can be carried 
out in order to investigate the feasibility of land treatment of such soil. 
Biodegradation potential of a particular hydrocarbon waste can be de- 
termined by the extensive chemical characterization of the petroleum- 
contaminated soil. Huesemann (1994b) has provided useful guidelines on 
carrying out laboratory feasibility studies on potential of land treatment of 
petroleum-contaminated soil. 

Laboratory mesocosms to study biodegradation of petroleum hydrocar- 
bons in contaminated soil can be prepared in open glass or metal trays as 
follows: 

1. Trays containing 5-10 kg of contaminated or spiked soil are prepared. 

2. Oil and grease or TPH content is determined and adjusted in the range 
of 5-7% by diluting with clean soil. 

3. To obtain optimal soil moisture content for the microbial activity, soil 
moisture is adjusted to between 50 and 80% of the field capacity (water- 
holding capacity), usually between 10 and 16 g of water per 100 g of dry 
soil. 

4. Adjust the pH to around 7.0 using lime, caustic soda, elemental sulfur 
or ammonium sulfate. 

5. The trays should be incubated at the optimum temperature range for 
microbial degradation of 25-35 °C. 

6. For each 100 kg of oil to be degraded, 1 kg of nitrogen and 0.2 kg of 
phosphorus should be added as nutrient fertilizer to obtain an oil:N:P 
ratio of 100:1:0.2. 

7. The duration of the biotreatability study depends on the overall ob- 
jective of the project. In general, it is recommended to run for 3-6 
months. 

8. Oil and grease or TPH content, moisture and pH should be periodically 
monitored. 

9. The soil should be lightly raked or mixed at 1-2-week intervals to 
provide proper aeration, mixing, and moisture control. 

10. The moisture content should be monitored at 1- or 2-week intervals and 
the soil sprayed with water to adjust to the optimum moisture content. 

Monitoring the disappearance of oil and grease or TPH, as well as mois- 
ture, pH, and nitrogen is important during the treatability studies. Total 



140 A. Singh et al. 

heterotrophic or hydrocarbon-degrading microbial counts may also be 
monitored to evaluate the biodegradation process. It is important to use 
the same sampling strategy and methods throughout the treatment period. 

5.2.7 
Composting 

While composting of yard and municipal wastes has been performed 
for decades, composting of hydrocarbon-contaminated soils represents 
an emerging ex-situ biological technology. Composting has been demon- 
strated to be effective in biodegrading explosives and polycyclic aromatic 
hydrocarbons (PAHs) in soils (USEPA 1996, 1998). In the composting of 
contaminated soil, organic amendments including manure, sewage sludge, 
compost, yard wastes, and food processing wastes are often added to supple- 
ment the amount of nutrients and readily degradable organic matter in soil. 
Sewage sludge and compost containing abundant nitrogen, organic mat- 
ter, and high microbial diversity, with total microbial populations higher 
than fertile soils, have great potential in bioremediation. A small-scale 
biotreatability method (Van Gestel et al. 2003) for composting technology 
is described here: 

1. Two insulated composting bins can be used, one filled with biowaste 
(vegetable, fruit, garden, and paper waste) only, and the other filled 
with a mixture of biowaste and petroleum-oil-contaminated soil at 
a 10:1 ratio (fresh mass). 

2. Dewatered sewage sludge or matured compost can be used instead of 
biowaste. 

3. Spruce bark can be used as a bulking agent at the ratio of soil to bulking 
agent, 1:3 on a volume basis. 

4. The soil should be collected from the top 15 cm of the soil surface and 
air dried and sieved to pass a 2-4 mm sieve. 

5. The soil can be spiked with commercial crude oil or diesel oil at a con- 
centration to obtain a concentration of 5-10 g/kg after mixing with the 
biowaste. 

6. The initial pH is adjusted about 7.0-7.4. 

7. The composting process is controlled using airflow and moisture con- 
tent. 

8. Aerobic composting can be performed for 12 weeks. 



5 Feasibility Studies for Microbial Remediation 141 

9. At regular time intervals, the content should be turned to avoid prefer- 
ential aeration pores. 

10. Compost samples for chemical and microbiological analyses should be 
taken every time the compost is mixed. 

11. Microbial counts, dry matter content, pH, temperature, electrical con- 
ductivity, and exhaust gas composition should be regularly monitored. 

12. Microbial composition of the biowaste-only composting bin serves as 
a reference for the composting process of contaminated soil. 

13. To investigate the degradation rate of oil in soil alone, a soil-only exper- 
iment (without organic amendments) should also be run as a control. 

Composting technologies can be applied to cleanse contaminated soil ex 
situ. By adding an organic matrix to contaminated soil the general microbial 
activity is enhanced and also the activity of specific degraders, which may 
be found in the contaminated soil or introduced along with the organic 
material. Biodegradation rates in composting systems have been found to 
be slightly higher than in land treatment of hydrocarbons and lower than 
in slurry reactors. 

5.2.8 
Scale-Up 

The data obtained from the small-scale biodegradation experiments can 
be used to design full-scale biotreatment systems. In most cases slurry 
bioreactors can be directly scaled up. The US EPA has suggested a three- 
tier approach before a full-scale application of the technology in the field 
(US EPA, 1991; McFarland et al. 1991): 

1. Laboratory screening to establish the occurrence and rate of biodegra- 
dation and establishing optimum process parameters 

2. Bench-scale testing to establish performance of the process parameters 
and cost estimate for the scale-up of appropriate technologies 

3. Pilot testing on the most promising technology to establish system design 
and detailed cost structure 

Land- or reactor-based full-scale bioremediation systems have been suc- 
cessfully used to clean up hydrocarbon-contaminated soils and sludges. 
More information on the scale-up of bioremediation technologies can be 
obtained in the literature (Huesemann 1994; Cutright 1995; Crawford and 
Crawford 1996; Loehr and Webster 1996; Von Fahnestock et al. 1998; Alle- 
man and Leeson 1999; Stegmann et al. 2001; Singh and Ward 2004). 



142 A. Singh et al. 

5.3 

Process Monitoring and Evaluation 

It is important to make sure that system operation and monitoring plans 
have been developed for the land treatment operation. Regular monitor- 
ing is necessary to ensure optimization of biodegradation rates, to track 
constituent concentration reductions, and to monitor vapor emissions, mi- 
gration of constituents into soils beneath the landfarm (if unlined), and 
groundwater quality. If appropriate, ensure that monitoring to determine 
compliance with storm water discharge or air quality permits is also pro- 
posed. 

1. Molecular composition of a petroleum contaminant can be useful in 
estimating the biodegradation potential of the contaminated soil. Gas 
chromatography (GC) analysis (Chapt. 3) may identify easily biodegrad- 
able compounds such as straight chain alkanes. GC analysis of various 
volatile (benzene, toluene, ethyl benzene, and xylenes) and semi- volatile 
(polynuclear aromatic hydrocarbons, PAHs) compounds are required 
by the regulatory agencies. However, gravimetric determination of oil 
and grease or TPH content following Soxhlet extraction can be used to 
design and optimize a reactor or land-based treatment process. 

2. Since abiotic processes such as dilution, adsorption, and volatilization 
can be responsible for hydrocarbon disappearance, criteria other than 
simple hydrocarbon disappearance should be used to assess biodegra- 
dation by microorganisms. Increase in the number of hydrocarbon- 
degrading bacteria as the bioremediation progresses provides evidence 
of biodegradation. Formation of colonies on the surface of a solidified 
mineral salts medium with silica gel, incubated in vapors of volatile 
hydrocarbons (Walker and Coleman 1976), can be used to enumerate 
hydrocarbon-degrading bacteria. Bacteria capable of degrading semi- 
volatile hydrocarbons (e.g., PAHs) can be enumerated by examining 
colonies on agar plates for their ability to visibly alter a layer of pre- 
cipitated insoluble hydrocarbon (Bogardt and Hemmingsen 1992). The 
modified most probable number (MPN) technique can be used for non- 
volatile hydrocarbons either by applying a floating sheen of oil to the 
surface of mineral medium or by placing hydrocarbons dissolved in 
a solvent in 24- or 96-well microtiter plates (Brown and Braddock 1990; 
Steiber et al. 1994; Haines et al. 1996). The presence of hydrocarbon- 
utilizing bacteria is detected by the emulsification or dispersion of sheen, 
by reduction of added iodonitrotetrazolium violet, or by the appearance 
of colored metabolites in the medium (see also Chapt. 13). 



5 Feasibility Studies for Microbial Remediation 143 

3. Since microbial communities play a significant role in biogeochemi- 
cal cycles, it is important to analyze the community structure and its 
changes during bioremediation processes (Chaps. 10 and 12). The tem- 
poral and spatial changes in bacterial populations and the diversity of 
the microbial community during bioremediation can be determined us- 
ing sophisticated molecular methods (van Elsas et al. 1998; Widada et al. 
2002). 

4. Biodegradation potential of a hydrocarbon-contaminated soil can be 
estimated by its chemical characterization and the relative biodegrad- 
ability of the contaminants. Mono aromatic compounds such as ben- 
zene and alkyl benzene and low molecular weight n-alkanes are eas- 
ily biodegradable as compared to high molecular weight and highly 
branched molecules. While PAHs with four or more rings are consid- 
ered recalcitrant, two or three ring PAHs can be degraded by different 
microbial species. 

5. The volatile constituents present in petroleum-contaminated soils tend 
to evaporate during biotreatment, particularly during tilling or plowing 
operations in land treatment and aeration of the bioreactors, rather 
than being biodegraded by bacteria. For compliance with air quality 
regulations, the volatile organic emissions should be estimated based on 
initial concentrations of the petroleum constituents present. Depending 
upon specific regulations for air emissions, control of VOC emissions 
may be required. Control involves capturing vapors and then passing 
them through an appropriate treatment process before being vented to 
the atmosphere. Control devices range from an erected structure such 
as a greenhouse or plastic tunnel to a simple cover such as a plastic sheet 
for land treatment and a carbon filter or biofQter for a slurry reactor. 

6. Solid-phase microextraction (SPME) has been used to monitor biodegra- 
dation of semivolatile hydrocarbons in diesel-fuel-contaminated water 
and soil (Eriksson et al. 1998) and of volatile hydrocarbons during bac- 
terial growth on crude oil (Van Hamme and Ward 2000). Although the 
method requires external calibration with several standard calibration 
curves, SPME was proven to be a rapid and accurate method for monitor- 
ing volatile and semivolatile hydrocarbons in petroleum biodegradation 
systems. 

5.4 
Bioaugmentation 

Bioaugmentation can be denned as the introduction of a large number 
of exogenous microorganisms into the environment of a biotreatment sys- 



144 A. Singh et al. 

tern. Diverse microorganisms, including many species of bacteria and fungi 
are known to degrade hydrocarbons. The most prevalent bacterial hy- 
drocarbon degraders belong to the genera Pseudomonas, Achromobacter, 
Flavobacterium, Rhodococcus, and Acinetobacter. Penicillium, Aspergillus, 
Fusarium, and Cladosporium are most frequently isolated hydrocarbon 
degrading filamentous fungi. Among the yeasts Candida, Rhodotorula, 
Aureobasidium, and Sporobolomyces are the hydrocarbon degraders most 
often reported (Van Hamme et al. 2003). Environmental and nutritional 
factors influence the presence, survival, or activity of microorganisms in 
contaminated soils. 

There are at least four different routes that result in the development of 
microbes capable of degradation of hydrocarbons at a certain site: 

1. The indigenous microflora are exposed to the contaminant long enough 
for genetic evolution to create a capacity to degrade the compound(s). 

2. The indigenous microflora, adapted to the local conditions, are exposed 
to one or more contaminating xenobiotic compounds. The bacteria ac- 
quire genes and degradation pathways from bacterial cells immigrating 
from elsewhere. 

3. The indigenous, well-adapted microflora are maintained ex-situ and 
then artificially supplied with the required degradative capacity. 

4. A bacterium that is thought to be competitive at the contaminated site is 
chosen. This may be a strain that is known to degrade the contaminant 
or one that is specifically constructed for this purpose. 

Bioaugmentation-related experiments can be conducted in slurry biore- 
actors described above. Bio augmentation studies can be carried out either 
using mixed cultures or individual pure strains. The effect of initial popula- 
tion size on biodegradation of contaminants can be determined by varying 
inoculum densities. The inoculum size can be varied from 10 5 to 10 9 CFU/g 
of soil in the bioaugmentation studies. The effect of a commercial or selec- 
tively developed inoculum on the rate of biodegradation, C0 2 evolution, 
time of lag phase after inoculation, and microbial population dynamics 
during biodegradation process can be monitored. 

5.5 

Effect of Surfactants 

The biodegradation rate of a contaminant depends on the rate of contam- 
inant bioavailability, uptake, and mass transfer. Bioavailability of a con- 
taminant in soil is influenced by a number of factors such as desorption, 



5 Feasibility Studies for Microbial Remediation 145 

diffusion, and dissolution. Use of chemical- or bio-surfactants in contam- 
inated soil can help overcome bioavailability problems and accelerate the 
biodegradation process. 

Biosurfactants, surface-active substances synthesized by living cells, 
have the properties of reducing surface tension, enhancing the emulsifi- 
cation of hydrocarbons, stabilizing emulsions, and solubilizing hydrocar- 
bon contaminants to increase their availability for microbial degradation. 
Biosurfactant-producing microbes play an important role in the acceler- 
ated bioremediation of hydrocarbon-contaminated sites (Rahman et al. 
2003; Shin et al. 2004). The low-molecular-weight biosurfactants (glycol- 
ipids, lipopeptides) are more effective than those of high molecular weight 
(amphipathic polysaccharides, proteins, lipopolysaccharides, lipoproteins) 
in lowering the interfacial and surface tensions (Mulligan 2005). 

Some simple laboratory experiments to study biosurfactant production 
and application in bioremediation are described here. 

5.5.1 

Screening of Microbial Cultures for Biosurfactant Production 

Different microbial cultures can be screened for biosurfactant production 
using the following method: 

1 . Prepare a series of 250-mL flasks containing 50 mL of sterile YPG medium 
(composition per L: 5 g peptone, 5 g yeast extract, 10 g glucose, pH 7.0) 
and incubate on a shaker (200 rpm) at 30 °C after inoculation with indi- 
vidual cultures. 

2. Add 1% glycerol after 24 h. 

3. Measure biomass content, biosurfactant production, surface tension, and 
emulsification activity at 12-24 h intervals. 

4. For biomass determination, filter the culture broth using GF/C filters, 
place the filters at 110 °C for 24 h, and weigh to calculate biomass (dry 
mass). 

5. Surface-active compounds can be extracted by liquid-liquid extraction 
using 10 mL of chloroform:methanol (2:1 ) mixture from 10 mL of the cell- 
free culture broth acidified with 1 N HC1 to pH 2. Concentrate the organic 
extracts by drying them overnight in a drying chamber at a temperature 
around 44 °C, and measure the mass of the biosurfactant. 

For purification of the biosurfactant to determine its properties and 
application, the culture broth is filtered through a centrifuge filter with 
lOkDa molecular weight cut-off at 6,000 g until the minimal amount of 



146 A. Singh et al. 

retentate is achieved. The retentate is diluted in 50% methanol in order 
to dissociate the micelles and filtered at 6000 g again. After collection of 
filtrate, methanol is evaporated under vacuum in a rotary evaporator at 
65 °C and the aqueous solution of the purified biosurfactant is lyophilized. 

Surface tension (mN/m) can be measured using a standard commercial 
tensiometer. The emulsification activity can be determined by adding a hy- 
drocarbon (xylene, benzene, n-hexane, kerosene, gasoline, diesel fuel, or 
crude oil) to the same volume of cell-free culture broth, vortexing for 2 min 
and letting stand for 24 h. The emulsification activity is determined as the 
percentage of height of the emulsified layer divided by the total height of 
the liquid column (Rahman et al. 2003). 

A blood agar lysis method can also be used for screening cultures for 
their biosurfactant-producing capabilities (Youssef et al. 2004). Culture is 
streaked onto blood agar plates and incubated for 48 h at 37 °C. The zones of 
clearing around the colonies indicate biosurfactant production. The diam- 
eter of the clear zones depends on the concentration of the biosurfactant. 

5.5.2 

Effect of Biosurfactants 

Biosurfactant preparations can be purchased from a commercial chemical 
supplier or purified from the culture broth as described above. For different 
hydrocarbons, a biosurfactant is added to the cultures to obtain concen- 
trations above and below the critical micelle concentration (CMC). The 
CMC value is determined by measuring surface tension in different dilu- 
tions of a 4 g/L solution of the biosurfactant. The value of CMC, expressed 
in mg/L, is obtained from the plot of the surface tension versus the loga- 
rithm of the concentration. A rhamnolipid biosurfactant concentration of 
50-2,000 mg/L is generally useful in biodegradation studies. 

The biodegradation experiment to study effect of biosurfactants can be 
conducted in 250 mL Erlenmeyer flasks containing 50 mL of the culture 
medium described before. Appropriate controls, such as no-biosurfactant 
and abiotic controls, are run along with the flasks containing different 
concentrations of the biosurfactant. Cultures are incubated on a shaker for 
7-14 days at 30 °C. 

5.5.3 

Effect of Chemical Surfactants 

Properties of chemical surfactants that influence their efficacy include 
charge (nonionic, anionic, or cationic), hydrophilic-lipophilic balance 
(HLB, a measure of surfactant lipophilicity), and CMC (the concentration 



5 Feasibility Studies for Microbial Remediation 147 

at which surface tension reaches a minimum and surfactant monomers ag- 
gregate into micelles). However, there is always a concern that the surfactant 
may get used preferentially as a carbon source instead of the contaminant. 
Hence, there is a need to provide a perspective as to when or how surfac- 
tants may be exploited in petroleum hydrocarbon degradation processes 
to improve rates and extents of degradation. 

Typical surfactant concentrations for washing of contaminant soil are 
1-2%, whereas the same contaminants may be solubilized in an aqueous 
solution at a surfactant concentration of 0.1-0.2%. Non-ionic surfactants 
within the HLB range of 11 to 15 can optimally support microbial degra- 
dation of hydrophobic contaminants. Nonylphenol ethoxylated surfactants 
with HLB 12 and 13 can substantially enhance biodegradation of hydro- 
carbons at surfactant concentrations greater than CMC value. Different 
groups of nonionic surfactants should be tested at different concentrations 
greater than their CMC during feasibility studies in soil microcosms or 
slurry reactors. 

5.6 

Optimization of Environmental Conditions 

The procedures described in the previous sections on different technologies 
can be used in studies of the factors affecting biodegradation rates and 
determining appropriate biotreatment strategy for contaminated soil. 

1. The optimum soil pH for hydrocarbon bioremediation in soil ranges 
from 6 to 8. Methods for adjusting pH usually include periodic appli- 
cation of lime and/or sulfur. The requirement of acid or alkaline solu- 
tions/solids for pH control is developed in biotreatability studies and 
the frequency of their application is modified during land treatment or 
slurry reactor operation as needed. In case of acidic soil (pH < 6), lime 
or calcium carbonate may be added to increase the pH to the required 
optimum range. For alkaline soil (pH > 8), elemental sulfur, ammonium 
sulfate, or aluminum sulfate may be added to lower the pH. 

2. Optimum temperature range for microbial degradation is 25 to 35 °C. 
Biodegradation rates are expected to slow considerably below 15 °C or 
above 40 °C. However, temperature cannot be maintained for land appli- 
cation. Land treatment of hydrocarbon-contaminated soils is difficult to 
operate in temperate and arid zones. Slurry bioreactors are always more 
useful in such places because environmental conditions can be more 
precisely maintained and with relative ease. 

3. During land treatment, soil microorganisms can only biodegrade petro- 
leum hydrocarbons within a limited range of favorable soil moisture 



148 A. Singh et al. 

conditions. If the soil is too dry, bacterial growth and metabolisms will 
be greatly reduced or even inhibited. Alternatively, if the soil is too wet or 
flooded, soil aeration will be greatly impaired which, in turn, will result 
in anaerobic conditions that are not conducive to hydrocarbon biodegra- 
dation. Since the moisture content at field capacity is strongly dependent 
on the soil type (clay and high organic matter soils retain comparatively 
higher moisture content), it is important to determine the moisture re- 
tention profile for each soil to be studied. The optimum moisture content 
for stimulating petroleum hydrocarbon biodegradation ranges from 50 
to 80% of the moisture content at field capacity. For example, if the soil 
moisture at field capacity was determined to be 20 g of water per 100 g of 
dry soil, the soil moisture content should be maintained between 10 and 
16 g of water per 100 g of dry soil. 

4. In order to limit the demand of oxygen by soil bacteria, it is important 
not to overload the soil with too high levels of oil contamination during 
land application. As outlined below, the optimum contaminant load- 
ing level for land treatment is about 5% (by weight) of oil. Maximum 
degradation rates are typically observed in the 10-15 cm upper plow 
layer if hydrocarbon concentrations are maintained around 5%. Addi- 
tion of peroxygen compounds may also help slowly release oxygen into 
the soil and thereby enhance the aerobic biodegradation of petroleum 
hydrocarbons. 

5. There are other processes such as volatilization, leaching, sorption and 
photo-oxidation that may cause the removal of certain hydrocarbon 
compounds or classes during biotreatment. It has been estimated that 
between 15 and 60% of fuel hydrocarbons (diesel, jet fuel, and heat- 
ing oil) can be lost during soil bioremediation by land treatment solely 
due to evaporation (Salanitro 2001). At room temperature (20 °C), most 
hydrocarbons with carbon numbers up to Q 5 or Ci6 readily evaporate 
from soil if in free contact with air. Even heavier hydrocarbons (> Ci 6 ) 
including three- and four-ring PAHs are likely to volatilize in intense 
sunshine. These competing loss mechanisms during field or laboratory 
bioremediation studies should be measured or estimated either by cal- 
culating a complete mass balance or by carrying out proper microbial 
control experiments. 



5.7 

Optimization of Nutritional Factors 

For biotreatment of petroleum hydrocarbons, bacteria that are both aerobic 
and heterotrophic are the most important in the biodegradation process. 



5 Feasibility Studies for Microbial Remediation 149 

Since microorganisms require organic and inorganic nutrients such as ni- 
trogen, phosphorus, magnesium, calcium, iron, and trace metals to support 
cell growth and sustain biodegradation processes, nutrients need to be sup- 
plemented during biotreatment in bioreactors or land in order to maintain 
active bacterial populations. However, excessive amounts of certain nutri- 
ents such as phosphate and sulfate can repress microbial metabolism. Im- 
portant nutrient sources for biotreatability or feasibility studies are shown 
Table 5.1. 

1. Nutrients are added for the growth and maintenance of microorgan- 
isms. By providing an appropriate balance of nutrients it is possible to 
achieve high level of growth of hydrocarbon-degrading bacteria and thus 
accelerated rates of hydrocarbon degradation. The typical non-carbon 
elemental composition of major bacterial components is nitrogen 12.5%; 
phosphorus 2.5%; potassium 2.5%; sodium 0.8%; sulphur 0.6%; calcium 
0.6%; magnesium 0.3%; copper 0.02%; manganese 0.01%, and iron 0.01% 
(Rehm 1993). Use of appropriate concentrations and ratios of nutrients 
can avoid a situation where growth is limited by depletion of one essential 
nutrient while all other nutrients may be present in excess. 

2. An oil carbon content of 80% can be assumed for the purpose of cal- 
culating C:N or C:P ratio. Although a wide range of C:N and C:P ratios 
has been recommended in the literature, an oil:N:P ratio of 100:1:0.2 
can be used for the feasibility studies. Thus, for each 100 kg of oil to be 
degraded, 1 kg of nitrogen and 0.2 kg of phosphorus can be added as 
nutrient fertilizer in the preliminary studies. Optimum C:N ratio can 

Table 5.1. Important nutrient sources for biotreatability or feasibility studies 

Nutrient source Examples 

Defined medium 

Nitrogen KN0 3 , NH4NO3, NH 4 C1 

Phosphorus KH2PO4, sodium tripolyphosphate (NasPaOio) 

Potassium KNO3 

Calcium CaCl 2 • 2H2O 

Magnesium MgS0 4 • 7H 2 

Iron FeCl 3 • 6H 2 

Trace metals MnS0 4 • H 2 0, CuS0 4 • 5H 2 0, ZnCl 3 • 4H 2 0, H3BO3, CoCl 2 • 6H 2 0, 

Na 2 Mo0 4 • 2H 2 

Complex medium 

Nitrogen Yeast extract, Peptone, Urea, NPK fertilizer 

Phosphorus NPK fertilizer 

Magnesium NPK fertilizer 

Trace metals Yeast extract, NPK fertilizer 



150 A. Singh etal. 

determined in microcosms or slurry bioreactors by varying C:N ratio 
from 10 to 100. 

3. For land treatment, nutrient supply methods usually include periodic 
application of solid fertilizers, while tilling to blend soils with the solid 
amendments, or applying liquid nutrients using a sprayer. For bioslurry 
reactors, a blend of solid nutrients can be added, which is quickly dis- 
solved in the medium due to continuous mixing. The composition of 
nutrients is developed in lab treatability studies. 

4. The inability of microbes to completely mineralize a contaminant and 
transform it to other organic compounds means that these organisms 
require other substrates to support their growth. The contaminants 
are transformed by "co-metabolic" processes, where a second substrate 
serves as primary energy or carbon source. 

5. Using a naturally selected and acclimated indigenous bacterial culture 
originating from the sludge is supplemented with a carefully designed 
blend of nutrients containing sources of nitrogen, phosphate, a complex 
protein, essential minerals, and a surfactant. The bioslurry reactor sys- 
tem can promote growth of a highly active microbial population and 
rapid conversion of the petroleum hydrocarbons at the rate of about 1% 
petroleum hydrocarbons degraded per day (Ward et al. 2003). 

6. Generally high molecular weight PAHs (five-ring) are only biodegraded 
in the presence of other hydrocarbons such as lower molecular weight 
PAHs or complex hydrocarbon mixtures such as crude oil. If these neces- 
sary co-substrates are absent, the co-metabolic biodegradation of higher 
molecular weight PAHs cannot proceed. 



5.8 
Conclusions 

Bioremediation is a cost effective and environmentally friendly hydrocar- 
bon-contaminated soil remediation technology. The successful bioreme- 
diation of contaminated soils depends on numerous environmental pa- 
rameters and operational factors, which need to be optimized in order to 
achieve maximum treatment benefits. Even under optimal conditions it is 
unlikely that all contaminants will be removed from the soil. This incom- 
plete biodegradation may be acceptable if the residual hydrocarbons can 
be shown to have no significant impact on ecological receptors and do not 
pose a risk to groundwater resources. 

The effectiveness of bioremediation depends on the success in identi- 
fying the rate-limiting factors and optimizing them in the feasibility and 



5 Feasibility Studies for Microbial Remediation 151 

biotreatability studies. Feasibility studies are essential and may have enor- 
mous impact on the cost of the full-scale operation. Depending on the 
site, nature of contamination, and type of soil, various methods for fea- 
sibility studies are currently available. These methods can be modified to 
accommodate the lab facilities and equipment availability. Sometimes it 
is difficult to extrapolate the results directly from the laboratory to the 
field. Nevertheless, successful bench- or pilot-scale test results are mostly 
useful in designing the full-scale bioprocessing system for bioremediation 
of hydrocarbon-contaminated soil. 

One of the main barriers to greater effective adoption of bioremediation 
technologies is the perception that the processes are very project-specific, 
requiring much customization. There is a need to develop more robust and 
technologically versatile processes that do not require significant research 
and development for each project (Ward 2004). Government funding initia- 
tives and the market favor use of more controlled and accelerated processes 
and that are typically more predictable (Srinivasan 2003). Hughes et al. 
(2000) have provided guidance with regard to selection of bioremediation 
configuration for treatment of different classes of chemicals. 

The choice of technology configuration based on application of such 
principles should precede the design of a feasibility study, and the latter 
then used to confirm and validate the effectiveness of the technology. 

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6 



Feasibility Studies forMicrobial Remediation 
of Metal-Contaminated Soil 

Franz Schinner, Thomas Klauser 



■ Introduction 

Objectives. Heavy metal contamination of soil is widespread due to metal 
processing industries, tannery, combustion of wood, coal and mineral oil, 
traffic, and plant protection. The toxic effects of heavy metals result mainly 
from the interaction of metals with proteins (enzymes) and inhibition of 
metabolic processes. In contrast to organic pollutants, metals are not min- 
eralized by microorganisms but can be oxidized or reduced, transformed 
to different redox stages, or complexed by organic metabolites. 

Besides excavation and deposition, a conventional treatment for decon- 
tamination of metal-polluted soil is extraction using mineral acids. The 
disadvantages of such a treatment are the destruction of soil, high costs 
of acids, and low acceptance. Alternative remediation strategies to reduce 
bioavailability of metals are: (1) immobilization with repeated addition of 
substances such as carbonate, phosphate, apatite, zeolite, clay minerals, 
peat, or humus: and (2) bioleaching with heterotrophic microorganisms, 
preferably fungi. The latter method represents a sustainable remediation 
treatment of metal-polluted soils (Wasay et al. 1998, Schinner et al. 2000). 
Autotrophic bacteria as known from ore leaching cannot be recommended 
for this treatment due to the high buffer capacity of soil compared to ore. 
The most limiting factor of heterotrophic leaching is the availability of 
inexpensive carbohydrates, such as molasses or the refuse from process- 
ing sugar, fruits, or white wine. Heterotrophic leaching can be done off 
site, on site, and in situ, and is an alternative treatment for the decon- 
tamination of metal- containing filter dusts (Schinner and Burgstaller 1989; 
Burgstaller et al. 1992) and industrial sludges. 

Principle. Metal-contaminated soil is supplemented with carbohydrates to 
increase the excretion of organic acids by autochthonous (procedure A) 
or inoculated (procedure B) fungi. Organic acids mobilize metals that are 
eluted from soil by percolation with water. 

Theory. In natural ecosystems fungi play an important role in the mobiliza- 
tion of nutrients and trace elements from soils. Autochthonous soil fungi 



Franz Schinner, Thomas Klauser: Institute of Microbiology, Leopold Franzens University, 
Technikerstrafte 25, 6020 Innsbruck, Austria, E-mail: franz.schinner@uibk.ac.at 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



156 R Schinner, T. Klauser 

but also inoculated single strains or mixed populations increase organic 
acid production after the addition of carbohydrates. Fungi mobilize met- 
als from mineral soils by excretion of acidifying protons (acidolysis), by 
excretion of organic acids forming cyclic organometal complexes (complex- 
olysis), or by redoxolysis with organic acids. Some of the Deuteromycetes, 
especially members of the genera Aspergillus and Penicillium, produce 
various organic acids, such as citric, oxalic, tartaric, gluconic, succinic, 
formic, and amino acids (Burgstaller and Schinner 1993; Gadd 1999). The 
metal- containing eluate obtained after percolation of soil with water can 
be regenerated by conventional metal recovery, such as precipitation, ion 
exchange, or biosorption. 

■ Equipment 

• Percolator: a filtration unit (funnel and suction flask; e.g., Nalgene, 
500 mL; Nalge Nunc, Naperville, IL, USA), with gauze mat (e.g., Schlei- 
cher & Schull TG 100; Schleicher & Schull, Dassel, Germany) instead of 
filter 

• Vacuum pump 

• Multichannel peristaltic pump 

• Timer (for circuit switching peristaltic pump and vacuum pump) 

• Atomic absorption spectrometry (AAS) or inductively- coupled plasma- 
atomic emission spectrometry (ICP-AES) for metal analyses 

• High performance liquid chromatography (HPLC) for analyses of or- 
ganic acids (optional): e.g., column AMINEX-HPX 87H (Bio-Rad, Her- 
cules, CA, USA); flow rate 0.6 mL/min; column temperature 41 °C, wave- 
length 210 nm, eluant 4 mM H 2 S0 4 (Womersley et al. 1985) 

■ Materials and Reagents 

• Quartz sand 

• Sawdust 

• Grain (e.g., barley, wheat, rye) 

• Organic-acid-producing fungi, e.g., Aspergillus sp. or Penicillium sp. 

• Complex substrate: e.g., dried refuse of sugar production, fruit, or white 
wine processing 

• Molasses solution: 150 g (< 75% dry mass)/L of water 



6 Feasibility Studies for Microbial Remediation of Metal- Contaminated Soil 157 

• Sterile KC1 solution: 10 g/L 

• Deionized water 

■ Sample Preparation 

Use air-dried, sieved (< 5 mm) soil. 

■ Procedure 

Procedure A: Metal Leaching with Autochthonous Microorganisms 

1. Assemble percolation units, pour about 10 mm quartz sand onto the 
gauze mat. Connect tubes to flasks containing molasses and water, and 
install pumps and timer. Prepare 3-4 replicates. 

2. Weigh 150 g of sieved soil, 15 g of sawdust, and 15 g of complex substrate 
into a beaker, mix, and pour it into the percolator. 

3. Add 60 mL of molasses solution, 50 mL of KC1 solution and 70 mL of 
water to rewet substrate. 

4. Start percolation with molasses solution, use the peristaltic pump at 
a flow rate of 20mL/h for 15 h every day. Repeat this percolation on 
days 2, 3, and 6. 

5. On days 3, 5, and 7, use water instead of molasses solution as percolation 
fluid. 

6. For the suction of the fluid substrate and water and for additional aera- 
tion, start the vacuum pump every 90 min for 25 min without interrupt- 
ing the percolation (from the beginning to the end of experiment). 

7. To measure the leaching efficiency, take daily samples for metal detection. 
After 15 h of percolation with molasses or water add 120 mL of water onto 
the soil, suck off each percolator for 5 min, centrifuge 5 mL of eluate for 
15 min at 10,000 g. The supernatant is used for quantification of metals 
and organic acids. Repeat this procedure every day in the same way. 

8. After 3 days add 15 g of complex substrate and mix it into the topsoil of 
each percolator. 

Procedure B: Metal Leaching with Bioaugmentation 

1. Prepare the inoculum for bioaugmentation as follows: 

1.1. Sterilize 30 g of grain together with 30 mL of water in a 500-mL 
Erlenmeyer flask for 35 min at 121 °C. 



158 R Schinner, T. Klauser 

1 .2. Inoculate the substrate with organic-acid-producing fungi. Use 5 mL 
of spore suspension or collect spores from a stock culture to which 
sterile ringer solution is added to wash away spores. 

1.3. Incubate the inoculated grain 7-10 days at 25 °C to produce spores. 

1.4. Add 200-300 mL of sterile KC1 solution and shake thoroughly. 
Transfer the spore-containing suspension to a sterile flask. About 
10 8 spores/mL are required. 

1.5. Store spore suspensions for the preparation of further inoculates at 
-20 °C. 

2. Perform the procedure for metal leaching as described for procedure A, 
except for step 3: To rewet and inoculate soil and substrate, add 70 mL 
of spore suspension instead of 70 mL of water to 60 mL of molasses 
solution and 50 mL of KC1 solution. After 3 days (step 8) add 20 mL of 
spore suspension together with 15 g complex substrate and mix it into 
the topsoil of each percolator. 

Monitoring Metal Remediation 

Determine the pH value (Chapt. 2), the heavy metal content (using AAS 
or ICP; Chapt. 3), and eventually organic acids in the eluate using HPLC 
(Womersley et al. 1985). Additionally, soil enzyme activity (Chapt. 17), 
microbial biomass (Chapt. 14), or fungal biomass of soil (Rossner 1996) 
can be analyzed. 

■ Calculation 

Calculate the amount of leached metals from the sum of metal contents 
measured each day, considering soil dry mass and extraction volume. 

■ Notes and Points to Watch 

• Metal-leaching experiments with soil percolation attain decontamina- 
tion rates of 40-80%. 

• Inoculation with organic-acid-producing strains can result in a higher 
leaching efficiency. 

• Mixed cultures of fungi are more efficient than single strains. 

• The clay fraction of soils contains more metals than the silt and sand 
fraction. 

• The leaching efficiency of soil microorganisms depends strongly on the 
soil buffer capacity. Acidification of soil (pH < 6) may be necessary. 



6 Feasibility Studies for Microbial Remediation of Metal- Contaminated Soil 159 

• The leaching efficiency of soil microorganisms depends on the quality 
and quantity of soil organic matter, and on the aeration of soil. 

• The duration of percolation and the addition of molasses and water must 
be optimized for each soil. 

• In situ bioleaching of metal-contaminated soils needs effective drainage 
systems. 

• Depending on the soil material needed for monitoring analyses, a larger 
volume of soil (for example, 300 g soil in a 1 L percolator) can be used. 

References 

Burgstaller W, Schinner F (1993) Leaching of metals with fungi. J Biotechnol 27:91-116 
Burgstaller W, Strafter H, Wobking H, Schinner F (1992) Solubilization of zinc from filter 

dust with Penicillium simplicissimum: Bioreactor leaching and stoichiometry. Environ 

Sci Technol 26:340-346 
Gadd G.M (1999) Fungal Production of citric and oxalic acid: Importance in metal speciation, 

physiology and biogeochemical processes. Adv Microb Phys 41:47-92 
Rossner H (1996) Fungal biomass by ergosterol content. In: Schinner F, Kandeler E, Ohlin- 

ger R, Margesin R (eds) Methods in soil biology. Springer, Berlin Heidelberg New York, 

pp 49-51 
Schinner F, Burgstaller W (1989) Extraction of zinc from industrial waste by a Penicillium 

sp. Appl Environ Microbiol 55:1 153-1 156 
Schinner F, Huber W, Atzwanger M, Klauser T (2000) Metodo per il trattamento di suoli 

inquinati da composti da metalli pesanti (Method for the treatment of soils polluted 

by heavy metal compounds and related plant of treatment). Patents: IT/01. 12.00/IT 

RM000631, IT/01. 12.00/IT RM000632 
Wasay S.A, Barrington S.F, Tokunaga S (1998) Using Aspergillus niger to bioremediate soil 

contaminated by heavy metals. Bioremediation J 2:183-190 
Womersley C, Drinkwater L, Crowe JH (1985) Separation of tricarboxylic acid cycle acids 

and other related organic acids in insect haemolymph by high-performance liquid 

chromatography. J Chromatogr 318:112-116 



7 Feasibility Studies for Phytoremediation 
' of Metal-Contaminated Soil 

Aleksandra Sas-Nowosielska, Rafal Kucharski, 
Eugeniusz Malkowski 



7.1 
Introduction 

Phytoremediation, which is the use of herbaceous plants and trees to stabi- 
lize, recover, or volatilize pollutants in contaminated soil, is considered an 
emerging new technology. The application of phytoremediation is said to 
be environmentally friendly, relatively low in cost, and high in public accep- 
tance. However, there are still a number of limitations that affect its imple- 
mentation on a large scale. The most considerable limitations are: narrow 
range of contaminant concentrations within which the method can be ap- 
plied (potential of plant toxicity), dependence on weather, time-dependent 
growing season, and requirement for management of by-products. Until 
recently, the most commonly applied phytoremediation methods have been 
phytoextraction and phytostabilization, particularly for soils polluted with 
heavy metals. 

Phytoremediation is more a biological than a technical approach, and 
it is difficult to create a definitive protocol that could be applied to any 
polluted site. The limiting factors differ from site to site, and therefore each 
project protocol, must be customized to site-specific conditions. 

7.2 
Phytoextraction 

Phytoextraction is a biological method that utilizes properties of specific 
species of plants to take up and accumulate pollutants from soil. Certain 
species, called hyperaccumulators, may accumulate metals up to several 
percent of their dry mass (Brooks 1998; McGrath et al. 2000). Unfortu- 
nately, the practical use of these plants for phytoextraction is limited due 



Aleksandra Sas-Nowosielska, Rafal Kucharski: Land Management Department, Insti- 
tute for Ecology of Industrial Areas, Kossutha 6 St, 40-833 Katowice, Poland, E-mail: 
sas@ietu.katowice.pl 

Eugeniusz Malkowski: Department of Plant Physiology, Faculty of Biology and Environ- 
mental Protection, University of Silesia, Jagiellonska 28 St, 40-032 Katowice, Poland 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



162 A. Sas-Nowosielska et al. 

to sparse production of biomass and problems with mechanical harvest- 
ing. Nevertheless, genetic research is being conducted to increase biomass 
production. 

A compromise to the problem of low accumulation properties is to use 
plant species with extensive biomass production to compensate for the 
lower metal accumulation rates. Such plants remove certain amounts of 
metals, but the process is very slow. The addition of chelators to con- 
taminated soil enhances metal uptake, an approach known as "induced 
phy to extraction" (Salt et al. 1998). Depending on local climate and chem- 
istry of pollutants being removed, the most commonly used species for 
heavy-metal extraction are Brassica and Helianthus. 

A schematic diagram of the processes in an induced phytoextraction 
project aimed at cleaning up soils moderately contaminated with lead, 
cadmium, and/or zinc is presented in Fig. 7.1. The description contains 
step-by-step procedures necessary to perform a phytoextraction project 
including theory, legal considerations, technical aspects, as well as logistic 
issues and equipment. 

7.2.1 
Treatability Study 

Site Characterization 

Site characterization includes the following information: 

• Site contaminants (targets: Pb, Zn, and/or Cd) 

• Existing vegetation (indicating potential for plant growth) 

• Proximity to water (for irrigation) 

• Proximity to electrical supply 

• Site accessibility for vehicles and farm equipment 

• Field observations 

• Historical site activities 

• Summary of regional hydrology/geology 

The purpose of a treatability study is to identify optimal conditions for 
metal uptake into the aboveground portion of the plants and to determine 
if the soil to be treated will support plant growth. The further objectives 
are to evaluate and select the appropriate plant species and soil amend- 
ments, and to optimize plant growth for maximal removal of metals from 
soils. Treatability studies include short-term investigations for evaluating 
the growth and metal uptake potential of selected plant species under con- 
trolled conditions in a growth chamber or greenhouse. 



7 Feasibility Studies for Phytoremediation of Metal-Contaminated Soil 



163 



CONTAMINATED SITE 
CHARACTERIZATION 



TREATABILITY STUDY 



STREAMLINE TEST 



SITE 
PREPARATION 



PLANTING 



BIOMASS 
PRODUCTION 



HARVESTING OF 
METAL-CONTAINING CROP 



SAFE CROP 
DISPOSAL 



ECONOMICAL 
CONSIDERATIONS 



RISK ASSESSMENT 



LEGAL CONSIDERATIONS 



PLANT CARE 
FERTILIZING 



METAL - MOBILIZING 
SOIL AMENDMENTS 



CROP 
PRE-TREATMENT 



Fig. 7.1. Induced phytoextraction process 



164 A. Sas-Nowosielska et al. 

A bridge between routine treatability studies and full-scale field appli- 
cations is the Streamline Test (Sas-Nowosielska et al. 2001), which in com- 
bination with a routine laboratory study allows for a rapid and inexpensive 
assessment of soil features across the entire site to be treated. 



Performance of Treatability Study 

1. Soil from the site being investigated should be collected from the top 
0-25 cm depth horizon and be well homogenized and sieved to pass 
a 4 mm sieve. The soil is placed inside 400-cm 3 plastic pots filled previ- 
ously with drainage of approx. 100 g of clean pea/river gravel (2-8 mm 
diameter) placed in the bottom. Each pot should be filled with 300 g of 
the sieved soil and watered with 100 mL of distilled water prior to plant- 
ing the seeds. The seeds should be placed on the surface of the soil in 
a circular pattern, covered with a thin layer of soil and moistened with 
an additional 40 mL of water. 

2. Pots are then placed inside the growth chamber or greenhouse on sep- 
arate plastic saucers in order to prevent leaching of metals and to avoid 
cross contamination. The seeds should be kept wet during germination 
to avoid additional compaction of the soil surface and exposure of seeds. 

3. Plants should be adequately fertilized 10 days after germination using 
commercial mixtures, and 14-16 days after germination the seedlings 
should be thinned as needed. 

4. Application of soil amendments should be completed approx. 1 week 
prior to harvest. Amendments should be administered in a single dose, 
or in three doses if necessary. 

Five replicates of each treatment are recommended. 



Sampling and Analytical Procedures 

An accurate chemical analysis of soil and plants by an accredited laboratory 
is a key requirement for conducting a successful treatability study. The 
results should be reliable, as they will indicate the effectiveness of the 
process and what changes maybe needed to enhance the phytoremediation 
process. For most of the sampling and analytical activities described in this 
chapter, ISO Standards are recommended (see References and Chapts. 1-3). 
Samples should be collected to determine the concentration (spatial 
variation) of the target metals at the site, initially and during the phytore- 
mediation activities. The actual number and location of samples should be 
based on the final layout of the field. Samples collected should be extracted 
and analyzed following the ISO Standards (see References). 



7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 165 

Plant material should be washed with tap water in an ultrasonic washer 
to remove soil particles and then dried at 70 °C. Approximately 1 g of dried 
ground material should be wet-ashed using concentrated nitric acid in 
a microwave system. Concentrations of metals should be analyzed by flame 
atomic absorption spectrophotometer (FAAS) or by inductively coupled 
plasma spectroscopy (ICP-AES). 

A chain of custody should be maintained during sampling activities and 
Quality Assurance/Quality Control procedures are to be followed. Analyt- 
ical work in phytoremediation projects focuses on soil and plant analyses, 
preceded by the sampling process. The related regulations are listed below. 
In the case of soil, the following information on the investigated material 
is required: 

• Soil texture (hydrometric method) 

• Soil pH in 1 N KC1, H 2 0, and 0.01 M CaCl 2 ; soil-to-solution ratio of 1:2.5 
(Chapt. 2) 

• Soil electroconductivity; soil to solution ratio of 1:2.5 

• Soil organic matter content by loss on ignition (Chapt. 2; Houba et al. 
1995) 

• Cation exchange capacity (CEC), according to ISO 13536 (1995) 

• Content of NO" and NH+ (Chapt. 2; Houba et al. 1995) 

• P content in water extract and in an ammonium lactate-acetic acid extract 
(Houba etal. 1995) 

• K content soluble in 8 M KC1 (Houba et al. 1995) 

• Amorphous Al and Fe content, using an ammonium oxalate -oxalic acid 
extract method (Houba et al. 1995), and obtaining the concentration of 
Al and Fe with ICP analysis (Houba et al. 1995) 

• Metal extraction and determination (optional): 

- Total heavy-metal and other major cation concentrations are deter- 
mined after extraction with aqua regia. Soil should be ground to pass 
a 0.25 mm sieve; concentrations of metals is analyzed by FAAS or 
ICP-AES (Chapt. 3). 

- Bioavailable fraction: 5 g of air-dried soil ground to < 0.25 mm is 
extracted with 50 mL of 0.01 M CaCl 2 for 5 h and the concentration of 
metals is analyzed in the extract using FAAS or ICP-AES. 

- Potentially available metal fraction: 4 g of air-dried soil ground to 
pass a < 2-mm sieve are extracted with 40 mL of 0.43 N HNO3 for 4 h. 



166 A. Sas-Nowosielska et al. 

The concentration of metals is analyzed in the extract using FA AS or 
ICP-AES. 

- Exchangeable cations: extraction is according to ISO 13536 (1995) as 
for CEC; concentration is measured by AAS. 

Streamline Test 

Site characterization and treatability studies currently are conducted se- 
quentially, prior to the initiation of full-scale planting. The purpose of 
these activities is to describe the nature and extent of contamination at 
the target site, and to determine if, and under what conditions, proposed 
plant species will extract the target contaminants. The present approach is 
time consuming, expensive, and may not lead to a successful scale-up for 
field scale application of phytoextraction. The traditional treatability study 
is conducted in greenhouse conditions with controlled air temperature, 
light, water regime, and homogenized soil. These carefully controlled con- 
ditions often do not mimic real world conditions. A streamline test (ST) 
is an attempt to combine the treatability study and site characterization 
into an integrated single effort (Sas-Nowosielska et al. 2001). The concept 
of the ST was based on a geostatistical assumption that an adequately dis- 
tributed number of soil samples may describe the distribution of metals 
across an investigated site. The variability of lead and cadmium contents 
in soil was estimated in previous field scale phytoextraction experiments 
(Kucharski et al. 1998). Based on these findings, it was assumed that two 
crossed strips covering approx. 20% of the total site surface would be suffi- 
cient to represent the entire area for site characterization purposes. Topsoil 
samples were taken outside and inside the strips, and analyzed for con- 
tents of metal. Comparison of average concentrations of lead, cadmium, 
and zinc in the soil inside and outside showed no significant differences. 
It was concluded that the ST better reflects the "real world" conditions as 
compared to the usual treatability study. The ST provides an early indica- 
tion/screening of the suitability of the site for the application of phytore- 
mediation. 



7.2.2 

Full-Scale Application 

Seedbed Preparation and Plant Protection 

Plant protection consists of applying herbicides for weed control and in- 
secticides to combat herbivore insects. The principle of application should 
follow the rules of good agriculture practice. Once the fertilizer and insec- 
ticides have been applied, the seedbed will be prepared for planting. 



7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 167 

The site preparation activities in general require the site to be cleared, 
cleaned and the soil developed into a condition that will allow planting. 
The following are examples of the kinds of obstacles that should be taken 
into consideration: 

- Quantity and extent of surface debris 

- Depth to water table 

- Potential for flooding from off the site 

- Location of depressions in the soil that will collect water and drown 
plants 

- Location of a reliable water source for irrigation 

Fertilization 

Most plant species have varying nutritional needs. Fertilization protocols 
should be prepared individually after a soil analysis indicating the required 
nutrients for the species used. An example of this was demonstrated when 
Brassica was used to clean lead-contaminated soil. Brassica sp. have lower 
phosphorus and potassium requirements than other species, but require 
abundant nitrogen and supplemental sulfur to promote rapid vegetative 
growth. Thus, in accord with the findings of the soil fertility analysis, the 
site had to be fertilized with nitrogen, phosphorus, potassium, and sulfur. 
Generally, other nutrients are not found to be deficient. Fertilizer type, 
placement, and quality play an important role in the success of the crop 
development. For example, Brassica sp. are very sensitive to salt damage 
from fertilizer placed too close to the seed or with the seed. 

Irrigation 

Irrigation is used to achieve maximum plant growth pertaining to soil 
moisture. The objective is to maintain the recommended soil moisture 
levels for each individual crop (depending on plant species and/or cultivar) 
during the project's first and second crops. The soil moisture has to be 
kept at the optimum level. According to local conditions, various irrigation 
systems can be used (dripping, overhead sprinkling, wand-style spraying). 
The initial irrigation after planting should wet the soil profile to a depth 
of 15 cm. Care should be taken to not apply too much water. Brassica sp. 
do not respond well to standing water. The soil should be kept damp but 
not saturated until the seedlings emerge. This may require irrigation every 
day for sandy soils and every 5-7 days for heavy soil types. The site should 
be checked daily to determine if the plants need irrigation. Timing of 
irrigations will depend on many variables, such as the size of the plants, 



168 A. Sas-Nowosielska et al. 

rainfall, temperature, soil type, and the rate of evapotranspiration, to name 
a few. A simple tool that can be used to aid a project's irrigation needs is 
a tensiometer. 

Soil Amendments 

• For Brassica sp.: K 3 EDTA (2.5 mmol/kg of soil; stock solution concentra- 
tion 50%) and acetic acid (5 mmol/kg of soil; stock solution concentration 
80%). 

• For Helianthus sp.: K 3 EDTA (5 mmol/kg of soil; stock solution concentra- 
tion 50%) and acetic acid (5 mmol/kg of soil; stock solution concentration 
80%). 

Doses of amendments should be calculated individually for each kind 
of soil and applied as a diluted solution through the irrigation system or 
tractor driven sprayer. 

Species Used for Phytoextraction 

• Brassica juncea, the species commonly used for lead phytoextraction, 
grows well on fertile, well-drained soils. Successful B. juncea establish- 
ment requires a fine, firm seedbed that is free of weeds and rubble. All 
vegetation that will compete with the phytoremediation crop should be 
removed. The objective is to produce a seedbed that will give the newly 
seeded crop maximum opportunity to germinate and grow. The addi- 
tion of high quality clean organic matter should be made only when it 
is essential. B. juncea has a low tolerance for high salinity and poorly 
aerated soils. A well-drained soil provides optimum conditions for rapid 
germination and uniform emergence. The seeding rate for B. juncea is 
15 kg/ha, and the ideal plant population is 1 10-160 plants/m 2 , which pro- 
duces 1.5-3.0 g seeds/m 2 . Planting depth will depend on soil moisture. 
The seeds must have good contact with moist soil to achieve maximum 
germination and emergence. Ideal planting depth is 1-2 cm. Under dry 
sandy conditions the depth may have to be adjusted to 2.5 cm, but plant- 
ing deeper than 2.5 cm can result in poor emergence and reduced plant 
population. 

• Other plant species, such as sunflower, can be used for a phytoextraction 
process. This plant can produce a high amount of biomass (60 t/ha), but 
heavy-metal accumulation is rather low (about 100 mg Pb/kg dry soil). 

• Some plants, termed "hyperaccumulators," take up toxic elements in sub- 
stantial amounts, resulting in concentrations in aboveground biomass 
over 100 times of those observed in conventional plants. It is econom- 
ically hard to grow plants hyperaccumulating toxic metals because of 



7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 169 

their very low biomass production and difficulty of harvesting (Blaylock 
etal. 1997). 



Harvesting 

All aboveground biomass should be removed at harvest at the right time 
of plant maturity, after adding the chelating amendment. Application of 
the soil amendment should be made in a manner so that as little of the 
amendment as possible contacts the plant. This can be done with a hand- 
held application wand that directs the liquid amendment at the soil below 
the plants leaves, or by using an automatic dispenser (Kucharski et al. 2000). 

The physical removal of the plants can be started 7-10 days after the 
application of the amendment. The plants should be cut as close to the 
ground as possible. This can be accomplished with a fodder harvester, 
which cuts plants into 3-5 cm pieces loaded onto the adjacent trailer. 

After harvest, the soil must be retilled and prepared for the next crop's 
planting. In general, the techniques for the first crop are used for the subse- 
quent crops, but less fertilizer should be used for them. The recommended 
rates must be determined. 

Crop Disposal 

Important steps after harvest are reduction of the crop volume and re- 
moval of excess water. These will improve technical parameters of harvested 
biomass in terms of further processing and reduce transport costs to the 
treatment or disposal site. Volume reduction of contaminated plant mate- 
rial can be achieved by composting, compaction, or pyrolysis processes. 

Composting and compaction should be considered as pre-treatment 
steps, since a large volume of contaminated biomass will still exist after 
both processes. Total dry-mass loss of contaminated plant biomass is an 
advantage of composting as a pretreatment step. It will reduce costs of 
transportation to a hazardous waste disposal facility and of deposition, 
or of transportation to other facilities where final crop disposal is to take 
place. Compaction does not result in total dry mass loss of plant biomass 
but works faster than composting. Pyrolysis is also considered a pretreat- 
ment step, since metal-contaminated material (coke breeze) is one of the 
end-products. Significant more volume/mass reduction of contaminated 
plant biomass is observed than from composting or compaction. More- 
over, pyrolytic gas is recovered in the process (Sas-Nowosielska et al. 2004). 

For final disposal, incineration of contaminated plant material in non- 
ferrous or cement rotary kilns or in a municipal waste incineration plant 
is considered the most promising method because it significantly reduces 
the biomass of harvested plant material. Deposition in hazardous waste 
disposal facilities seems to be the simplest way to dispose of contaminated 



170 A. Sas-Nowosielska et al. 

crops. However, it is not completely adequate since significant amount of 
heavy-metal contaminated material will remain in the environment and 
costs of its disposal are high (Sas-Nowosielska et al. 2004). 

Monitoring 

Routine monitoring should include collection and analysis of the following: 
soil, airborne deposition, plants, vadose zone moisture, ground water, and 
irrigation water. In addition, routine soil chemistry and weather monitoring 
should be conducted. Weekly visits to the site are recommended to examine 
the growth of plants, soil moisture, appearance of pests, etc. 

7.2.3 
Conclusions 

The following remarks attempt to summarize the up-to-date observations 
concerning phytoextraction: 

• The method is applicable to cleansing sites contaminated to a moderate 
or medium degree. 

• Although described in literature as a lead-extraction method, significant 
amounts of zinc and cadmium can thus be extracted. High concentra- 
tions of zinc in soil can impair plant growth. 

• Results of laboratory and bench studies should be cautiously transferred 
to the field; conclusions carelessly drawn from experiments on pollutant- 
spiked soil can be a source of very serious errors. 

• Each new clean-up project has to be custom-tailored due to significantly 
varying soil conditions and pollutant distribution. 

• Induced phytoextraction, where costs of reagents contribute to the major 
portion of all project expenditures, should for economic reasons only be 
used on particularly valuable areas. 

• Continuous phytoextraction, which may be considered a type of natural 
contaminant attenuation by selected species of plants, is applicable to 
the sites where time is not a driving factor. 

• It is highly recommended to use indigenous species for phytoextraction, 
as they are considerable cheaper than exotic species, and do not create 
adaptation problems. 

• Although EDTA, a commonly used metal- chelating agent, was found not 
to exert any adverse effects on soil bacterial and fungal life, it should be 
very carefully applied considering its potential for mobilization of metal 
into other compartments of the environment. 



7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 171 

7.3 

Phytostabilization Potential for Soils 

Highly Contaminated with Lead, Cadmium and Zinc 

The use of certain plant species to immobilize contaminants in the soil 
and ground water through accumulation and absorption by roots, adsorp- 
tion onto roots' epidermis, or precipitation within the root zone is called 
phytostabilization. This term further infers a physical stabilization of soil. 
Phytostabilization does not remove contaminants from the soil, but reduces 
the hazards to human health and environment. 

Plants are used to cover the soil surface to prevent erosion, reduce 
water percolation, serve as a barrier to prevent direct contact with the 
soil-immobilized contaminants, and to control soil pH, gases, and redox 
conditions (Vangronsveld et al. 1995). Plant roots may change soil pH by 
release of exudates or through the production of C0 2 during root respira- 
tion. 

Phytostabilization is a site stabilization technique that reduces the risk 
of soil contaminants through the use of soil amendments that induce the 
formation of insoluble contaminant species. The method essentially con- 
sists of a combination of the use of immobilizing soil additives to reduce 
the bioavailability of heavy metals in contaminated soil and the creation 
of a dense vegetation cover. Bioavailability as a function of remediation 
treatment can be quantified based on the contaminant enrichment factor 
(EF), which is the ratio of contaminant metal concentration in plant tissue 
to the total concentration in the soil. 

7.3.1 

Evaluation of Site Contaminants 

The investigator must establish a procedure, using less aggressive extrac- 
tion, for evaluating the level of contaminant initially associated with the 
solid phase. The relative extractability of the contaminant of interest is 
then evaluated before and after a given stabilization treatment. In essence, 
such techniques are an abbreviated sequential extraction and subjected to 
the same empirical limitations with respect to interpretation as is the full 
procedure. 

Sampling Setup 

The first sampling takes place before the technology is started. All contam- 
inated areas should be sampled by using the procedures described in ISO 
Standards. The following general soil parameters are quantified: 



172 A. Sas-Nowosielska et al. 

• Soil texture (hydrometric method) 

• Soil pH in 1 N KC1, H 2 and 0.01 M CaCl 2 ; soil to solution ratio of 1:2.5 
(Chapt. 2) 

• Soil electroconductivity; soil to solution ratio of 1:2.5 

• Soil organic matter content by loss on ignition (Chapt. 2; Houba et al. 
1995); 

• Cation exchange capacity (CEC), according to ISO 13536 (1995); 

• Content of NO" and NH+ (Chapt. 2; Houba et al. 1995); 

• P content in water extract and in an ammonium lactate-acetic acid extract 
(Houba etal. 1995); 

• K content soluble in 8 M KC1 (Houba et al. 1995); 

• Amorphous Al and Fe content, by an ammonium oxalate-oxalic acid 
extract method (Houba et al. 1995), t concentrations of Al and Fe being 
analyzed with ICP (Houba et al. 1995) 

• Metal extraction and determination (optional; see Sect. 7.2.1) 

7.3.2 

Logistic Considerations 

As in case of phytoextraction, the following is to be examined before the 
technical activity in the project is started: 

• Existing vegetation (indicating potential for plant growth) 

• Proximity to water (for installation of irrigation system) 

• Site accessibility for vehicles and farm equipment 

• Field observations 

• Historical site activities 

• Summary of regional hydrology/geology 

7.3.3 
Additives 

Soil amendments for phytostabilization should inactivate metal contam- 
inants, reducing their bioavailability, and preventing leaching and plant 
uptake. Some amendments, e.g., phosphate fertilizer, have secondary ben- 
efits such as supplying plant nutrients. 



7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 173 

Limestone and organic materials (Li and Chaney 1998), natural and 
synthetic zeolites, phosphate minerals (apatite, calcium phosphate, am- 
monium polyphosphate, etc.), iron and manganese oxides, are suggested 
for metal stabilization (Knox et al. 2000, 2001) 

The application rates of the additives generally range from 0.5-5% by 
soil mass. The largest amounts of additives are generally needed for highly 
contaminated areas with high a percentage of bioavailable forms of con- 
taminant. 



7.3.4 
Plants 

Desirable features of species used for land phytostabilization are as follows 
(Berti et al. 1998; Vangronsveld and Cunningham 1998): 

- Tolerance to high concentrations of pollutant (Li and Chaney 1998) 

- Ability to create a dense root mat 

- Ability to accumulate pollutants in a non-edible underground part 

- Low maintenance requirements (watering, pest and weed control) 

- Resistance to the local climatic extremes 

In the course of investigation of very highly metal-polluted areas it has 
been found that Deschampsia caespitosa ecotype Warynski, a weed growing 
spontaneously on the soil close to zinc smelter dumps, creates very dense 
and durable plant cover upon being supplied with necessary nutrients. This 
ecotype has appeared to be relatively strong and healthy, in spite of poor soil 
conditions and inadequate watering. Species such as Brassica juncea, some 
cultivars of grasses (Agrostis tenuis, Festuca rubra) and some cultivars of 
hybrid poplars are also suggested for this purpose. 

7.3.5 

Full-Scale Application 

The methodology presented has focused on highly metal- contaminated 
areas with poor plant cover. Chemostabilization combined with phytosta- 
bilization is meant to prevent pollutant migration via wind, water erosion, 
and leaching. 

Agronomic input includes the nutrients necessary for vigorous growth 
of vegetation and rhizosphere microbes. It should be done based on a local 
Good Agriculture Practice. Before the soil contaminants are to be stabi- 



174 A. Sas-Nowosielska et al. 

lized, inorganic (nitrogen, phosphorus, potassium), and organic fertilizers 
(manure, compost, etc.) have to be applied. 

For using 5% superphosphate, lime at the rate of about 12 t/ha should be 
applied and mixed to a depth of 20 cm. After liming, the amendment should 
be mixed with the upper 10-cm layer of soil. Two weeks after amendment 
application, about 60 kg/ha of D. caespitosa seeds should be planted. This 
concentration eliminates creation of tufts and increases density of plant 
cover. 

7.3.6 

Effectiveness of Technology 

Phytochemostabilization can be achieved by creation of appropriate plant 
cover in combination with soil amendments. The approach we suggest 
is suitable for areas heavily polluted with bivalent heavy metals, where 
commercially available species for revegetation cannot survive. The overall 
goal is complete coverage of the contaminated surface with a plant canopy, 
whose growth is enhanced by chemicals, and which will simultaneously 
immobilize pollutants and continue to support plant growth. 

In particularly complex deterioration of soil, where chemical soil dam- 
ages are followed by its mechanical destruction, the concerted action of 
phyto- and chemostabilization may yield positive results. Evaluation of 
hazard reduction must be made to validate the effectiveness of phytostabi- 
lization. 

7.3.7 
Monitoring 

Phytostabilization does not remove contaminants from the soil. There are 
two objectives of monitoring during a phytostabilization process: 

• To evaluate the long term effectiveness of immobilization in revegetated 
areas, which leads to an estimation of the reduction of the long term 
leaching potential and the influence of vegetation on leaching risks 

• To evaluate plant uptake potential in relation to potential transfer of 
heavy metals into the food chain 

Monitoring the fate of contaminants, as well as the presence of additives 
over a long period of time is required, particularly in areas with strong 
impact from acid rain and possible changes in soil redox potential. Immo- 
bilization effectiveness and plant uptake determinations should be carried 
out during the growing seasons of several years. 



7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 175 

7.3.8 
Conclusions 

The combined chemo-phytostabilization method has the following advan- 
tages: 

• Phosphate as used in the method decreases the concentration of bivalent 
heavy metals in roots and shoots, and their bioavailable fraction in 
leachates, and also improves plant cover density. 

• Further, phosphate thus introduced in soil may facilitate the propagation 
of Deschampsia in the third year of growth by enhancing production of 
seeds, which germinate on bare soil between the tufts. 

• The procedure supports the growth of the root system and makes it 
stronger, resulting in increases of up to 70% water retention and reduced 
metal migration. 

• The growth of D. caespitosa is improved in the process at the expense of 
the growth rate of Cardaminopsis sp. This is a positive phenomenon, be- 
cause high heavy-metal accumulation rates in Cardaminopsis sp. shoots 
results in a potential introduction of heavy metals into the food chain. 

• Metal migration to lower soil levels is decreased by the procedure as 
a result of metal-chemical binding and the development of a strong plant 
cover. 

• An optimization study to evaluate phosphorus addition to the soil and 
satisfactory plant growth remains to be done, and the price of the additive 
is also a matter of concern. 

• Phosphate used as a fertilizer for metal contaminated soils in very high 
concentration is considered disadvantageous as it causes saturation with 
phosphate in the upper soil layers. This can lead to phosphate leaching. 
Phosphate use is therefore limited to areas with a deep water table where 
groundwater pollution by phosphate is unlikely, and where the greater 
benefit of obtaining healthy plant cover is unlikely to be achieved. 

• Phosphate is not recommended for arsenic-polluted soils, as competition 
between arsenate and phosphate can provoke increased arsenic levels in 
plants, causing risks of food-chain propagation and accumulation. 



Acknowledgements. The authors wish to express their thanks to Mr. Laymon 
Gray of Florida State University for his editorial contribution to this paper. 



176 A. Sas-Nowosielska et al. 

References 

Berti WR, Cunningham SD, Cooper EM (1998) Case studies in the field - in-place inac- 
tivation and phytorestoration of Pb-contaminated sites. In: Vangronsveld J and Cun- 
ningham SD (eds) Metal-contaminated soils: in situ inactivation and phytorestoration. 
Springer- Verlag, Berlin Heidelberg and RG Landes Co, Georgetown, TX, USA, pp 235- 
248 

Blaylock MJ, Salt DE, Dushenkov S, Zakharova O, Gussman C, Kapulnik Y, Ensley BD, 
Raskin I (1997) Enhanced accumulation of Pb in Indian mustard by soil-applied chelat- 
ing agents. Environ Sci Technol 31:860-865 

Brooks RR (1998) Phytochemistry of hyperaccumulators. In: Brooks RR (ed) Plants that 
hyperaccumulate heavy metals. Cab International, Wallingford, Oxon, UK, pp 15-53 

Houba VJG, Van der Lee J J, Novozamsky I (1995) Soil analysis procedures, other procedures 
(Soil and plant analysis, Part 5b). Dept Soil Sci Plant Nutr, Wageningen Agricultural 
University, pp 217 

ISO 11265 (1994) Soil quality - Determination of the specific electric conductivity 

ISO 11464 (1993) Soil quality - Pretreatment of samples for physico-chemical analyses 

ISO 13536 (1995) Soil quality - Determination of the potential cation exchange capacity and 
exchangeable cations using barium chloride solution buffered at pH = 8.1 

ISO 7888 (1985) Water quality - Determination of electrical conductivity 

ISO/CD/10381-5 (1995) Soil quality - Sampling 

ISO/DIS 10390 (1993) Soil quality - Determination of pH 

ISO/DIS 11047 (1994) Soil quality - Determination of cadmium, chromium, cobalt, copper, 
lead, manganese, nickel and zinc. Flame and electromatic thermal atomic absorption 
spectrometric methods 

ISO/DIS 11466 (1995) Soil quality - Extraction of trace metals and heavy metals soluble in 
aqua regia 

Knox AS, Seaman J, Adriano DC, Pierzynski G (2000) Chemophytostabilization of metals in 
contaminated soils. In: Wise DL, Trantolo DJ, Cichon EJ, Inyang HI, Stottmeister U (eds) 
Bioremediation of contaminated soils. Marcel Dekker, Inc, New York, Basel, pp 811- 
836 

Knox AS, Seaman JC, Mench MJ, Vangronsveld J (2001) Remediation of metal- and 
radionuclides-contaminated soils by in situ stabilization techniques. In: Iskandar IK 
(ed) Environmental restoration of metal-contaminated soils. Lewis Publ, Boca Raton, 
London, New York, Washington, DC, pp 21-60 

Kucharski R, Sas-Nowosielska A, Dushenkov S, Kuperberg JM, Pogrzeba M, Malkowski E 
(1998) Technology of phytoextraction of lead and cadmium in Poland. Problems and 
achievements. In: Symposium Proceedings, Warsaw'98, Fourth Int Symposium and 
Exhibition on Environmental Contamination in Central and Eastern Europe, pp 55 

Kucharski R, Sas-Nowosielska A, Krynski K (2000) Amendment application technology 
for phytoextraction. In: Symposium Program, Prague 2000, Fifth Int Symposium and 
Exhibition on Environmental Contamination in Central and Eastern Europe, Abstract, 
p376 

Kucharski R, Sas-Nowosielska A, Kuperberg M, Bocian A (2004) Survey and assessment. 
How urbanization and industries influence water quality. In: Integrated watershed man- 
agement - ecohydrology & phytotechnology, manual. UN Educational, Scientific and 
Cultural Organization, Venice, Italy, pp 45-60 

Li YM, Chaney L (1998) Case studies in the field - industrial sites: phytostabilization of zinc 
smelter- contaminated sites: the Palmerton case. In: Vangronsveld J, Cunningham SD 
(eds) Metal-contaminated soils: in situ inactivation and phytorestoration. Springer- 
Verlag Berlin Heidelberg, and RG Landes Co, Georgetown, TX, USA, pp 197-210 



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McGrath SP, Dunham SJ, Correl RL (2000) Potential for phytoextraction of zinc and cadmium 
from soils using hyperaccumulator plants. In: Terry N, Banuelos G (eds) Phytoremedi- 
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Salt DE, Smith RD, Raskin I (1998) Phytoremediation. Annu Rev Plant Physiol Plant Mol 
Biol 49:643-668 

Sas-Nowosielska A, Kucharski R, Korcz M, Kuperberg M, Malkowski E (2001) Optimizing 
of land characterization for phytoextraction of heavy metals. In: Gworek B, Mocek A 
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Sas-Nowosielska A, Kucharski R, Malkowski E, Pogrzeba M, Kuperberg M, Krynski K (2004) 
Phytoextraction crop disposal - an unsolved problem. Environ Pollution 128:373-379 

Vangronsveld J, Cunningham SD (1998) Introduction to the concept. In: Vangronsveld J, 
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Vangronsveld J, Van Assche F, Clijsters H (1995) Reclamation of a bare industrial area 
contaminated by non-ferrous metals: in situ metal immobilization and revegetation. 
Environ Pollution 87:51-59 



8 



Quantification of Hydrocarbon 
Biodegradation Using Internal Markers 

Roger C. Prince, Gregory S. Douglas 



■ Introduction 

Objectives. Soil contamination is invariably heterogeneous, and monitor- 
ing the loss of contaminant during bioremediation is often frustrated by 
this heterogeneity. But if the initial source of contamination was relatively 
homogeneous, it is possible to identify biodegradation as the selective loss 
of the most biodegradable components, while more recalcitrant molecules 
are conserved. Measuring the concentrations of a series of compounds us- 
ing gas chromatography (GC) coupled with mass spectrometry (MS), often 
in the selected ion monitoring (SIM) mode, allows this to be achieved with 
high precision. 

Hopanes have proven to be useful conserved internal markers for fol- 
lowing the biodegradation of crude oil contamination (Prince at al. 1994), 
trimethylphenanthrenes for following the biodegradation of diesel fuel 
(Douglas et al. 1992), and 2,2,3,3-tetramethylbutane and 1,1,3-trimethyl- 
cyclopentane for following the anaerobic biodegradation of gasoline and 
condensate (Townsend et al. 2004). Undoubtedly, there are many other 
compounds that could be used. Even if the "conserved" internal marker 
is itself eventually degraded, this will have the effect of underestimating 
the extent of biodegradation of compounds referred to it, making the ap- 
proach a conservative one. The principal requirements are that the samples 
under consideration initially had the same contaminant, and that the com- 
pound chosen as the "conserved" internal standard be amongst the least 
degradable in the mixture under study, and be present at a high enough 
concentration to be measured with good precision. 

Principle. Depending on the type of contamination, which can be deter- 
mined from the hydrocarbons present (Stout et al. 2002), the least biode- 
graded sample is identified, and candidate conserved species are identified. 
The ratios of various analytes to these species are then followed over time, 
and biodegradation is identified from their coherent loss. The concentra- 
tion of the conserved species (e.g., hopane) on an oil-weight basis may 

Roger C. Prince: ExxonMobil Research and Engineering Co., Annandale, New Jersey 08801, 
USA, E-mail: Roger.C.Prince@ExxonMobil.com 

Gregory S. Douglas: NewFields Environmental Forensic Practice LLC, Rockland, Mas- 
sachusetts 02370, USA 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



180 R.C. Prince, G.S. Douglas 

also be used to estimate the total quantity of oil that has been degraded 
(Douglas et al. 1994) within a sample. 

Theory. The biodegradation of hydrocarbons has been studied for al- 
most a century, and the overall process is quite well understood (Prince 
2002). Under aerobic conditions, n-alkanes and simply substituted mono- 
aromatic species are amongst the most readily biodegraded hydrocarbons, 
followed by the iso- and monocyclic alkanes, benzene and the simply alky- 
lated two and three-ring aromatics (Solano-Serena et al. 1999). More highly 
alkylated species, four- ring and larger aromatics (Douglas et al. 1994), and 
compounds containing tertiary carbons are more resistant to biodegrada- 
tion (Prince et al. 1994). Similar patterns are seen under methanogenic and 
sulfate-reducing conditions, with the apparent distinction that some cyclic 
alkanes are very readily degraded under these conditions (Townsend et al. 
2004). The biodegradation of at least some hydrocarbons, e.g., toluene, 
occurs under other anaerobic conditions as well (Chakraborty and Coates 
2004). 

Inevitably some analyte in any complex mixture is its least biodegrad- 
able compound. Referring the concentrations of other analytes to this com- 
pound provides a ready index of the extent of biodegradation of that analyte, 
and removes much of the variability in the absolute concentration of the an- 
alyte in soil and sediment samples. This is shown graphically in the figures. 
Figure 8.1 shows the biodegradation of 2-methylhexane over 100 days in 
samples from a condensate-contaminated anaerobic aquifer amended with 
a small amount of gasoline and incubated under sulfate-reducing condi- 
tions (Townsend et al. 2004). The raw data are seen in Fig. 8.1 A, the data 
referred to 1,1,3-trimethylcyclohexane as a conserved internal marker in 
Fig. 8. IB. Similarly, Fig. 8.2 shows the biodegradation of the sum of the 
USEPA priority pollutant polycyclic aromatic hydrocarbons (PAHs; Keith 
and Telliard 1979) in a historically contaminated refinery soil over a time 
span of 1.5 years (Prince et al. 1997). The raw data are seen in Fig. 8.2 A, the 
data referred to 17a(H),21/?(H)-hopane as a conserved internal marker in 
Fig. 8.2B. In both cases, the biodegradation of the target compound(s) is 
much more apparent in the B panels. 



■ Procedure 

The precise recipes for extracting and analyzing samples will depend on 
many site-specific variables, and we give only a broad description of the pro- 
tocols involved. Measurements made for regulatory compliance are usually 
specifically mandated by the regulators involved, and we do not discuss 
them here. Rather we focus on measurements made to assess whether 
biodegradation is proceeding, and whether bioremediation protocols are 



8 Quantification of Hydrocarbon Biodegradation Using Internal Markers 



181 



100 ■ 



arb. 
units 



20 ■ 








100 



°/c 



■ 



40 80 

Days 



120 



20 ■ 








40 



Days 



80 



120 



Fig. 8.1. A The biodegradation of 2-methylhexane under sulfate-reducing conditions in sam- 
ples collected from a condensate-contaminated aquifer, amended with 1 uL of gasoline (per 
50 g sediment, 75 mL groundwater) and incubated in the laboratory under sulfate-reducing 
conditions (Townsend et al. 2004). The individual incubations were carefully assembled 
with equal weights of sieved sediments in each bottle, yet the raw data are still very hetero- 
geneous. B The data and referenced to the concentration of 1,1,3-trimethylcyclohexane in 
each sample 



140 



arb. 
units 



20 





200 400 

Days 




200 400 

Days 



600 



Fig. 8.2. Biodegradation of the 16 USEPA Priority Pollutant PAHs in a refinery soil. The data 
(the sum of the concentrations) were collected after a bioremediation protocol of adding 
slow release nutrients was initiated (Prince et al. 1997). A Although the soil was tilled during 
the treatment, and individual samples were sieved prior to analysis, the raw data are still 
very heterogeneous. B The data referenced to the concentration of 17a(H),21/?(H)-hopane 
in each sample 



indeed stimulating the process. This is best done by comparing samples 
from a site undergoing active bioremediation with samples from a similarly 
contaminated site with no intervention. Unfortunately, this is often impos- 
sible, and samples collected during active bioremediation protocols have 
to be compared with samples taken at the beginning of the remediation. In 
either case, absolute amounts of contaminants in "replicate" samples are 
likely to be log-normally distributed (Limpert et al. 2001 ), and changes due 



182 R.C. Prince, G.S. Douglas 

to biodegradation will be difficult to detect unless the conserved-marker 
approach is used. 

Sample Preparation 

Sample preparation is fundamentally different if the compounds of concern 
are in the gasoline or diesel and higher range. For soils, sediments, and water 
samples contaminated with gasoline, the appropriate extraction procedure 
is "purge-and-trap" analysis (Uhler et al. 2003). For soils contaminated with 
kerosene, diesel, heating, or crude oil it is more appropriate to extract the 
hydrocarbons into a solvent and inject the solvent-hydrocarbon mixture 
directly into the GC (Douglas et al. 1992, 2004). 

Internal Standards 

Often it is appropriate to add surrogate internal standards prior to extrac- 
tion. These may be added for two fundamentally distinct reasons. One is 
to assess the efficiency of the extraction protocol: fluorobenzene is often 
used for "purge-and-trap" analyses, while o-terphenyl is often used in sol- 
vent extractions. The second is to add compounds to check that the mass 
spectrometer is working correctly: deuterated compounds are often used 
(Uhler et al. 2003; Douglas et al. 1992, 1994, 2004). 

"Purge-and-Trap" 

"Purge-and-trap" protocols for the extraction of volatile hydrocarbons 
are described in USEPA methods 5030B: "Purge-and-Trap for Aqueous 
Samples," and 5035: "Closed-System Purge-and-Trap and Extraction for 
Volatile Organics in Soil and Waste" (USEPA 2003). Although the technical 
aspects are discussed in the EPA Method, the target analytes to which this 
method is applied includes only eight hydrocarbons present in gasoline 
(benzene, toluene, ethylbenzene, m-, p-, and o-xylene, styrene, and naph- 
thalene). This is inadequate for detailed characterization of gasoline and 
other light hydrocarbon products and for measuring conserved species. Uh- 
ler et al. (2003) have modified Method 8260 to quantitatively measure more 
than 100 diagnostic gasoline-related compounds ranging from isopentane 
to dodecane in nonaqueous phase liquid products, water, and soil. Due 
to the wide range of solubilities and volatilities of these compounds (e.g., 
benzene versus dodecane), caution must be exercised when analyzing these 
additional compounds by the purge-and-trap methods and careful calibra- 
tion and monitoring of analyte-recovery efficiencies should be performed 
(Uhler et al. 2003). 

In essence, an appropriate amount of sample to give a response within 
the calibrated range of the GC system is flushed (purged) with an inert gas 
to transfer the analytes of interest to a trap. When the purging is complete, 



8 Quantification of Hydrocarbon Biodegradation Using Internal Markers 183 

which usually takes several minutes, the trap is rapidly heated to transfer 
the sample into the GC column. If the sample is a soil sample, sufficient 
clean water is added prior to the purging to make a fluid slurry. The initial 
sampling must be done rapidly and into tightly sealed vessels to prevent 
any loss of volatile components during sample collection and storage. In 
our hands, samples containing about 1 pi of gasoline are appropriate for 
analysis (Townsend et al. 2004). 

Solvent Extraction 

Solvent extraction protocols are described in USEPA method 3500B: "Or- 
ganic extraction and sample preparation" (USEPA 2003). Soil or sediment 
samples are dried by mixing them with enough anhydrous sodium sul- 
fate to make a freely flowing dry mixture. Typical samples may require an 
equal weight of sodium sulfate, and it is important to mix thoroughly and 
for some time (perhaps 20 min) to allow the drying agent to hydrate and 
dry the sample. Samples are then serially extracted, at least three times, 
with an appropriate solvent (e.g., methylene chloride or methylene chlo- 
ride/acetone 1 + 1), perhaps in a Soxhlet extraction device, by accelerated 
solvent extraction (ASE), or by supercritical fluids. 

The extracts are dried with sodium sulfate, filtered, and then concen- 
trated as appropriate. It is important that this solvent-evaporation be done 
carefully to minimize the loss of lighter volatile components, such as the 
two-ring aromatics. Only in rare cases where it is known that there are no 
volatile compounds should it be allowed to proceed to dryness. Automated 
devices are available, but solvent-evaporation can be done manually under 
a gentle stream of dry nitrogen gas at ambient temperature. 

Depending on the minimum detection limits required (Douglas et al. 
2004), and the presence of interfering compounds, it maybe appropriate to 
process the solvent extract on an alumina or silica column to isolate "clean" 
fractions of saturate, aromatic, and polar compounds. This is described in 
detail in USEPA method 361 1: "Alumina column cleanup and separation of 
petroleum wastes" and USEPA method 3630 "Silica Gel Cleanup" (USEPA 
2003). Often the two hydrocarbon fractions (saturate and aromatic hy- 
drocarbons) are combined, concentrated to an appropriate volume, and 
amended with additional internal standards to allow quantitation; again 
deuterated compounds are often used. In our hands, 1 p.L injections of 
samples containing about 5 mg of crude oil/mL solvent are appropriate for 
analysis (Douglas et al. 1992, 2004). 

Gas Chromatography and Mass Spectrometry (GC/MS) 

This requires an appropriate high-resolution capillary column equipped 
with a mass spectrometer (McMaster and McMaster 1998; Hubschmann 



184 R.C. Prince, G.S. Douglas 

2000). USEPA methods 8260 and 8270D (USEPA 2003) provide GC/MS 
protocols for the measurement of volatile and semi-volatile hydrocar- 
bons, respectively. As noted above, the EPA protocols are not designed 
for petroleum product analysis and have been modified by various inves- 
tigators to increase the number of petroleum-specific target compounds 
(Douglas and Uhler 1993; Uhler et al. 2003) and improve the sensitivity of 
the methods (Douglas et al. 1994, 2004). 

For the modified EPA Method 8260 (Uhler et al. 2003) compounds are 
identified and quantified using full-scan mass spectrometry (typically from 
m/z = 35-300) for the extended volatile hydrocarbon target analyte list (109 
gasoline-specific compounds). The advantage of full-scan analysis is that 
additional compounds can always be evaluated, and extracted ion plots of 
compound classes (e.g., alkylcyclohexanes, Townsend et al. 2004) can be 
obtained to determine that the products are derived from the same source. 
Although the full-scan GC/MS approach is not as sensitive as selected ion 
monitoring (SIM), it is generally adequate for volatile hydrocarbon analysis. 

In contrast, it is essential to use selected ion monitoring (SIM) in the 
modified EPA Method 8270 (Douglas et al. 1992, 2004). This protocol al- 
lows the measurement of the major paraffins and isoparaffins, the aro- 
matics on the USEPA list of priority pollutants (Keith and Telliard, 1979) 
and their alkylated forms, and the steranes and hopanes that are so valu- 
able in discriminating different crude oils (Peters et al. 2004). The most 
significant modifications of the USEPA Method are the inclusions of the 
dibenzothiophenes, alkylated PAHs, steranes and hopanes that provide 
petroleum source identification and bioremediation efficacy information 
(Douglas et al. 2002). 

Analytes are identified by the retention times of authentic standard com- 
pounds, and by reference to mass spectral libraries such as those distributed 
by NIST/EPA/NIH (NIST 2004). It is always appropriate to use more than 
one ion to identify analytes in the initial samples to assess whether there 
are any interfering species present, and if so, how to account for them. 

For research purposes it is usually possible to arrange the concentra- 
tions of analytes to fall into the linear range of detectability, which should 
be determined with a range of calibration standards. A lot of work has gone 
into optimizing detection limits for the analysis of complex environmental 
samples for forensic applications (Douglas et al. 2004), but only the simplest 
precautions are needed for most studies quantifying biodegradation. Cer- 
tainly the mass spectrometer should be tuned with an appropriate standard, 
such as decafluorotriphenylphosphine, before every batch of samples, and 
standard samples and blanks should be included in every group of samples. 
Of course, if the analytical variability is large then the ability to detect an 
impact of a bioremediation protocol is reduced. Therefore, it is preferable 
to measure all the samples for a particular study at one time, or at least to 



8 Quantification of Hydrocarbon Biodegradation Using Internal Markers 185 

include control and reference samples with every batch. This may require 
that early samples be preserved until analysis; careful freezing or acidifica- 
tion to pH 2 with HC1 both work well. Furthermore, it is appropriate to set 
some "quality control" values that the standard samples must satisfy before 
the data are considered suitable for analysis. Guidelines for suitable control 
values are given in USEPA method 8270D (USEPA 2003) and in Page et al. 
(1995). 

■ Calculation 

We can calculate the percent of an analyte remaining (Figs. 8.1 and 8.2) 
from the equation: 

(Aq/Cs) 
% Remaining = — — - x 100 (8.1) 

8 (Ao/Q) 

A s concentration of the target analyte in the sample 

Cs concentration of the conserved compound in the sample 

A concentration of the target analyte in the initial sample 

C concentration of the conserved compound in the sample 

Alternatively the percent depletion of biodegradable analytes within the 
oil (Fig. 8.3) can be calculated using the equation: 

(Aq/Cq) - (As/Cs) .... 

%Loss = x 100 (8.2) 

(Ao/Co) 

Note that these equations work equally well in absolute concentration 
terms, or in arbitrary units, as long as the latter are obtained under identical 
conditions for all samples. 



Notes and Points to Watch 

The approach outlined here relies on the initial source of contamination 
being reasonably homogeneous. This is readily achieved in laboratory 
studies, and often pertains to acute contamination accidents such as oil 
spills. But chronic contamination may prove too heterogeneous for this 
approach to work without subdividing areas under consideration (e.g., 
Prince et al. 1997). For example, the composition of gasoline has changed 
over the years as more effective refinery processes have been introduced, 
and as the molecular composition has come under regulatory oversight. 



186 



R.C. Prince, G.S. Douglas 



Similarly, contamination at town gas sites and refineries may be from 
a mixture of sources. It is thus essential to take enough samples of the 
contamination prior to any remediation activities to delineate areas of 
similar and distinctly different contamination. 

It is important to minimize evaporative losses prior to analysis. This 
means carefully sealed sample vials for "purge-and-trap" analyses, and 
care during evaporative solvent removal from extracts. Including appro- 
priate surrogate compounds in the analysis can assess such losses. 

Biochemical intuition and published work will help identify potential 
analytes to be used as conserved internal compounds. Consistently neg- 
ative values for the % depletion of other analytes with respect to the 
"conserved" one will indicate that the "conserved" compound is in fact 
more degradable than the other analytes, and allow selection of a better 
standard compound (e.g., see Fig. 8.3) 

The simple analysis of Figs. 8.1 and 8.2 may be all that is needed to 
demonstrate that biodegradation is occurring, but more complicated 
models for biodegradation, taking into account the amount of oil, its 



100 



% 



depletion 



-100 




Referred to C 3 -phenanthrenes 
Referred to hopane 



/7C 1( 



nC 



34 



Co-phenanthrenes 



hopane 



pristane 

phytane 



naphthalene 



C-fluorenes 



chrysene 

C 3 -chrysenes 



nC 



20 



phenanthrene 



benzo[b]fluoranthene 



Fig. 8.3. Percent depletion plot for some alkanes, PAHs, and hopane in a degraded 
Alaskan North Slope crude oil (Douglas et al. 1994). The hatched series repre- 
sents the percent depletion of each analyte based on the C3-phenanthrenes (the 
trimethyl, methyl- ethyl, propyl and isopropylphenanthrenes) as the conserved inter- 
nal marker. Note that some compounds have a negative apparent depletion, indi- 
cating that the C3-phenanthrenes are less conserved than those analytes. The solid 
series represents the percent depletion based on the more biodegradation resistant 
17a(H),21^(H)-hopane. (Prince et al. 1994) 



8 Quantification of Hydrocarbon Biodegradation Using Internal Markers 187 

prior weathering, and the amount of available fertilizer, have been used to 
demonstrate the effectiveness of bioremediation in the field (Bragg et al. 
1994). 

• Biodegradation can be identified by the loss of biodegradable com- 
pounds, as discussed above. The loss of photochemically labile species 
can also be followed (Garrett et al. 1998; Douglas et al. 2002), as can the 
loss following extensive washing and evaporation (Douglas et al. 2002; 
Prince et al. 2002) and the increase of pyrogenic compounds following 
partial oil combustion (Garrett et al. 2000). Providing a sample of the 
initially spilled oil is available, these environmental processes can then 
be identified in samples collected from historical spills (Prince et al. 
2003). 

• The general approach can also be used to follow the biodegradation of 
any complex mixture of contaminants, such as polychlorinated biphenyls 
(Abramowicz 1995). 

References 

Abramowicz DA (1995) Aerobic and anaerobic PCB biodegradation in the environment. 

Environ Health Perspect 103 Suppl 5:97-99 
Bragg JR, Prince RC, Harner EJ, Atlas RM (1994) Effectiveness of bioremediation for the 

Exxon Valdez oil spill. Nature 368:413-418 
Chakraborty R, Coates JD (2004) Anaerobic degradation of monoaromatic hydrocarbons. 

Appl. Microbiol. Biotechnol. 64:437-446 
Douglas GS, Burns WA, Bence AE, Page DS, Boehm P (2004) Optimizing detection limits for 

the analysis of petroleum hydrocarbons in complex environmental samples. Environ 

Sci Technol 38:3958-3964 
Douglas GS, McCarthy KJ, Dahlen DT, Seavey JA, Steinhauer WG, Prince RC, Elmendorf DL 

(1992) The use of hydrocarbon analyses for environmental assessment and remediation. 

J Soil Contam 1:197-216 
Douglas GS, Owens EH, Hardenstine J, Prince RC (2002) The OSSAII pipeline spill: the 

character and weathering of the spilled oil. Spill Sci Technol Bull 7:135-148 
Douglas GS, Prince RC, Butler EL, Steinhauer WG (1994) The use of internal chemical 

indicators in petroleum and refined products to evaluate the extent of biodegradation. 

In: Hinchee RE, Alleman BC, Hoeppel RE, Miller RN (eds) Hydrocarbon remediation. 

Lewis Publ, Boca Raton, FL, pp 219-236 
Douglas GS, Uhler AD (1993) Optimizing EPA methods for petroleum contaminated site 

assessments. Environ Test Anal 2:46-53 
Garrett RM, Guenette CC, Haith CE, Prince RC (2000) Pyrogenic polycyclic aromatic hy- 
drocarbons in oil burn residues. Environ Sci Technol 34:1934-1937 
Garrett RM, Pickering IJ, Haith CE, Prince RC (1998) Photooxidation of crude oils. Environ. 

Sci. Technol. 32:3719-3723 
Hubschmann, H.-J. (2000) Handbook of GC/MS: fundamentals and applications. Wiley- 

VCH, Weinheim, Germany 
Keith LH, Telliard WA, (1979) Priority pollutants I. - a perspective view. Environ Sci Technol 

13:416-423 



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Limpert E, Stahel WA, Abbt M (2001) Log- normal distributions across the sciences: Keys 

and clues. Bioscience 51:341-352 
McMaster M, McMaster C (1998) GC/MS: A practical user's guide. Wiley- VCH, New York 
NIST (2004) NIST/EPA/NIH mass spectral library www.nist.gov/srd/mslist.htm 
Page DS, Boehm PD, Douglas GS, Bence AE (1995) Identification of hydrocarbon sources in 

benthic sediments of Prince William Sound and the Gulf of Alaska following the Exxon 

Valdez oil spill. In: Wells PG, Butler JN, Hughes JS (eds) Exxon oil spill: Fate and effects 

in Alaskan waters, ASTM Special Technical Publication #1219, American Society for 

Testing and Materials, Philadelphia, pp 41-83 
Peters KE, Walters CC, Moldowan JM (2004) The Biomarker guide, biomarkers and isotopes 

in petroleum exploration and earth history, vol 1-2, 2nd edn. Cambridge Univ Press, 

New York 
Prince RC (2002) Biodegradation of petroleum and other hydrocarbons. In: Bitton G (ed) 

Encyclopedia of environmental microbiology. Wiley, New York, pp 2402-2416 
Prince RC, Drake EN, Madden PC, Douglas GS (1997) Biodegradation of polycyclic aromatic 

hydrocarbons in a historically contaminated site, in: Alleman BC, Leeson A (eds) In situ 

and on-site bioremediation 2. Battelle Press, Columbus, OH, pp 205-210 
Prince RC, Elmendorf DL, Lute JR, Hsu CS, Haith CE, Senius JD, Dechert GJ, Douglas GS, 

Butler EL (1994) 17a(H),21/?(H)-hopane as a conserved internal marker for estimating 

the biodegradation of crude oil. Environ Sci Technol 28:142-145 
Prince RC, Garrett RM, Bare RE, Grossman MJ, Townsend GT, Suflita JM, Lee K, Owens EH, 

Sergy GA, Braddock JF, Lindstrom JE, Lessard RR (2003) The roles of photooxidation 

and biodegradation in long-term weathering of crude and heavy fuel oils. Spill Sci 

Technol Bull 8:145-156 
Prince RC, Stibrany RT, Hardenstine J, Douglas GS, Owens EH (2002) Aqueous vapor ex- 
traction: a previously unrecognized weathering process affecting oil spills in vigorously 

aerated water. Environ Sci Technol 36:2822-2825 
Solano-Serena F, Marchal R, Ropars M, Lebeault JM, Vandecasteele JP ( 1 999) Biodegradation 

of gasoline: kinetics, mass balance, and fate of individual hydrocarbons. J Appl Microbiol 

86:1008-1016 
Stout SA, Uhler AD, McCarthy KJ, Emsbo-Mattingly S (2002) Chemical fingerprinting of 

hydrocarbon. In: Murphy B, Morrison R (eds) Introduction to environmental forensics. 

Academic Press, New York, pp 135-260 
Townsend GT, Prince RC, Suflita JM (2004) Anaerobic biodegradation of alicyclic con- 
stituents of gasoline and natural gas condensate by bacteria from an anoxic aquifer. 

FEMS Microbiol Ecol 49:129-135 
Uhler RM, Healey EM, McCarthy KJ, Uhler AD, Stout, SA (2003) Molecular fingerprinting 

of gasoline by a modified EPA 8260 gas chromatography-mass spectrometry method. 

Int J Environ Anal Chem 83:1-20 
USEPA (2003) Index to EPA test methods, http://www.epa.gov/epahome/index/ 



9 



Assessment of Hydrocarbon Biodegradation 
Potential Using Radiorespirometry 

Jon E. Lindstrom, Joan E Braddock 



■ Introduction 

Objectives. Following environmental exposure to petroleum, acclimation 
of microbial communities to hydrocarbon metabolism may occur through 
selective enrichment of member populations possessing hydrocarbon cata- 
bolic pathways, induction or repression of enzymes, or genetic mutations 
resulting in new metabolic capabilities (Leahy and Colwell 1990). Measure- 
ments of carbon substrate mineralization in vitro can be used to assess the 
hydrocarbon biodegradative potential of microbial communities in envi- 
ronmental samples previously exposed to oil contamination in situ (Walker 
and Colwell 1976; Lindstrom et al. 1991; B0rresen et al. 2003). 

Using 14 C-labeled hydrocarbon substrates, mineralization of specific hy- 
drocarbon compounds can be tracked, and low levels of mineralization 
activity are detectable if sufficiently high specific activity substrates are 
employed. Model compounds can indicate the degree of a community's 
acclimation to various hydrocarbon classes (e.g., hexadecane for linear 
alkanes, toluene for monoaromatic hydrocarbons, or phenanthrene for 
polycyclic aromatic hydrocarbons (PAHs; Bauer and Capone 1988). By 
appropriately manipulating experimental conditions, this method may be 
used to assess the prior exposure of environmental samples to hydrocarbon 
contamination (Braddock et al. 1996; Braddock et al. 2003), or the effects of 
fertilization or other field treatments used to enhance in situ hydrocarbon 
degradation (Lindstrom et al. 1991). In addition, manipulation of nutri- 
ent levels or other amendments in the assay may be used in bench-scale 
treatability studies prior to initiating field-scale bioremediation efforts. 

Principle. A 14 C-labeled hydrocarbon substrate is added to a soil sample 
suspended in sterile diluent contained in a sealed volatile organic anal- 
ysis (VOA) vial. The sample is incubated under appropriate conditions 
(dictated by the experimental question), and microbial metabolism of the 
added substrate is measured by recovery of 14 C-labeled C0 2 evolved during 



Jon E. Lindstrom: Shannon & Wilson, Inc., 2355 Hill Road, Fairbanks, Alaska 99709, USA, 
E-mail: JEL@shanwil.com 

Joan F. Braddock: College of Natural Science and Mathematics, University of Alaska Fair- 
banks, Fairbanks, Alaska 99775, USA 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



190 J.E. Lindstrom, J.R Braddock 

incubation. Microbial activity is halted by adding a strong base at the end 
of the incubation period, which sequesters the C0 2 generated by microbial 
substrate mineralization as carbonates in solution. The 14 C-labeled C0 2 
is subsequently recovered by acidifying the suspension, then stripping the 
C0 2 from solution with nitrogen gas, and capturing it in a basic scintilla- 
tion cocktail. The 14 C0 2 derived from mineralization of the added labeled 
substrate is counted by liquid scintillation, and its radioactivity compared 
to that added with the labeled substrate. 

Theory. Petroleum is a complex mixture of hydrocarbons, and nitrogen-, 
sulfur- and oxygen-containing organic compounds; and the hydrocarbon 
fraction itself may be composed of hundreds of aliphatic, alicyclic, and 
aromatic compounds (National Research Council 1985). Heterotrophic 
biodegradation of the organic substrates in petroleum therefore occurs 
via a diversity of pathways, with metabolic intermediates funneled to cen- 
tral metabolic pathways leading to the production of microbial biomass 
and carbon dioxide (Wackett and Hershberger 2001). The fate of carbon in 
the substrate metabolized varies depending on the organism, the pathways 
used, and other factors. For example, biomass incorporation of glucose 
was approximately twice that of phenolic compounds in taiga forest floor 
samples, while respiration of C0 2 in these samples was significantly higher 
for phenolic compounds (Sugai and Schimel 1993). Despite the variation in 
carbon allocation among substrates and microbial communities, respira- 
tion of carbon dioxide is useful for monitoring biodegradation of organic 
substrates, particularly when the source of the carbon may be tracked by 
radioactive labeling. 

The protocol described here assesses the respiration activity of organ- 
isms in environmental samples. The procedure is designed to minimize the 
many factors affecting the actual mineralization activity in situ, except for 
the in situ microbial biomass and its potential to biodegrade the hydrocar- 
bons tested. The rate of 14 C0 2 production (r*, Bq/day) from a radiolabeled 
substrate is a function of the overall rate of C0 2 production (R) and the 
specific activity of the added label (Brown et al. 1991): 

r* = x R (9.1) 

(Sn+A) 

A* radioactivity of the labeled substrate added to the sample (Bq/g soil) 

S n in situ substrate concentration (|^g/g soil) 

A concentration of substrate added with the radiolabeled substrate (|ig/g 
soil) 

R rate of C0 2 production (|^g/day) from carbon sources in the sample 



9 Assessment of Hydrocarbon Biodegradation 191 

By adding to the sample an amount of the tested substrate (A) that is 
large compared to S n> the value of r* will mainly depend on A, rather than S n 
(Brown et al. 1991). As the amount of substrate added to the sample must be 
greater than the in situ concentration, and conditions in vitro are designed 
to minimize the various other factors affecting in situ mineralization rates, 
the value of r* reflects the microbial community's biodegradation potential 
only and is not a measure of in situ mineralization rates. 

The choice of incubation conditions may be used to assess the degree of 
a microbial community's acclimation to a given hydrocarbon substrate in 
the environmental sample, evaluate the effectiveness of field treatments, or 
establish optimum growth conditions for the community being studied. As 
the in situ mineralization rate may be attenuated due to nutrient deficien- 
cies or other environmental factors, radiorespirometric assays conducted 
with added nutrients or other amendments are useful for assessing the 
degree of community acclimation (suggesting prior exposure; Braddock 
et al. 1996; Braddock et al. 2003) to the hydrocarbon substrate or class of 
substrates (e.g., alkanes, monoaromatics, PAHs) being tested, since such 
environmental limitations are removed. 

A lag period following substrate addition is observed in the assay, with its 
duration commonly varying as a function of the solubility and molecular 
structure of the substrate (Brown et al. 1991). To measure the activity of 
the extant biomass present in the sample on collection, an appropriate 
incubation period must be chosen that is short enough to avoid in vitro 
acclimation of the native biomass to the added substrate, but long enough 
to detect its mineralization (see below). 



Equipment 

Incubators equilibrated to temperatures dictated by experimental re- 
quirements 

Apparatus for collecting C0 2 evolved from the soil suspension follow- 
ing incubation and liquid scintillation counter to detect the radioac- 
tivity associated with mineralization of the added labeled substrate. 
[A schematic of an apparatus suitable for stripping and capturing C0 2 
evolved from the soil suspension is shown in Fig. 9.1: Nitrogen gas is 
bubbled through the acidified soil suspension via a spinal needle (10- 
cm, 18-gauge deflected-point, non-coring, septum-penetrating needle 
with standard hub and stainless steel cannula; Popper and Sons, New 
Hyde, NY, USA) that pierces the silicone septum of the VOA vial. The gas 
stream strips the C0 2 from the suspension, and is conveyed to a Harvey 
trap (R.J. Harvey Instruments, Hillsdale, NJ, USA) containing acidified 
toluene via Tygon tubing attached to a 1-mL syringe sleeve cut to fit in 



192 



J.E. Lindstrom, J.R Braddock 



Gas flow direction 



Source of N 




40-mL VOA vial 
containing acidified 
soil suspension 



Harvey trap containing toluene 
acidified with 12 N HCI 



20-mL scintillation vial 
containing C0 2 -sorbing 
scintillation cocktail 



Fig. 9.1. Schematic diagram of stripping apparatus used to collect 14 C02 from samples 
following incubation. Nitrogen gas is bubbled through the sample, and the gas stream flows 
through a Harvey trap containing acidified toluene to trap any volatile hydrocarbons in 
the gas stream. Finally, 14 C02 is collected in a vial containing a C02-sorbing scintillation 
cocktail 



the tubing and equipped with a 16-gauge needle that pierces the VOA 
vial septum. The gas stream is bubbled through the acidified toluene in 
the Harvey trap to capture any labeled organic substrate that may have 
been stripped from the soil suspension. The gas stream containing the 
labeled C0 2 is then conveyed to a 20-mL scintillation vial fitted with 
a two-hole rubber stopper and glass tubing (a 1-mL glass pipette cut to 
a 5 cm length works well here for the glass tubing, as it provides a tapered 
and polished tip). The influent gas stream is bubbled through a 10 mL 
scintillation cocktail containing /?-phenylethylamine (PEA) to capture 
the C0 2 . Following a 15-min stripping period, the gas flow is stopped, 
the rubber stopper removed, and the scintillation vial capped and placed 
in a scintillation counter to determine the amount of recovered radioac- 
tivity. The stripping apparatus may be modified so that a number of 
samples may be run simultaneously. This requires a manifold equipped 
with valves and multiple sets of the apparatus described above. A single 
nitrogen tank can be connected to the manifold and used to strip 14 C0 2 
evolved from several soil suspensions in parallel.] 



9 Assessment of Hydrocarbon Biodegradation 193 

• Sterile and pre-cleaned or combusted 40 mL borosilicate VOA vials 
equipped with Teflon-lined, 0.125-mm-thick, silicone septa (e.g., I-Chem 
Brand; Nalge Nunc, Rochester, NY, USA) 

• Sterile 10-mL pipettes 

• 100-|iL syringe (Hamilton, Reno, NV, USA) 

• Syringes fitted with an 18-gauge needle 

■ Reagents 

• Sterile diluent: modified Bushnell-Haas broth (mineral nutrient; from 
Atlas 1993, but modified to contain l/10th strength FeCl 3 ) or Ringer's 
solution (Collins et al. 1989) 

• Hydrocarbon test substrate: Prepare a solution of non-labeled hydro- 
carbon substrate (hexadecane, benzene, phenanthrene, etc.) in acetone 
(2 g/L). Then add 14 C-labeled hydrocarbon substrate with sufficient spe- 
cific activity to obtain a final radioactivity of about 20 Bq/pL. 

• Toluene, acidified by adding HC1: Approximately 5-mL aliquots of toluene 
are used in the Harvey trap of the stripping apparatus (Fig. 9.1); add 
0.1 mL of 12 N HC1 to 5 mL of toluene placed in the trap. 

• Scintillation cocktail (Cytoscint ES; MP Biomedicals, Irvine, CA, USA) 
containing PEA to sorb C0 2 . Add 2.5 mL PEA to 7.5 mL Cytoscint and 
shake to mix; the PEA cocktail needs to be mixed within about 1 h of use. 

• ION NaOH to terminate incubation, and sequester evolved 14 C0 2 in 
solution 

• 12 N HC1 to release 14 C0 2 for recovery and counting 

■ Sample Preparation 

Use fresh soil samples. If samples must be stored, refrigerate them following 
collection. Sieve soil samples (2-mm mesh) to homogenize. 

■ Procedure 

Assay Preparation 

Soil samples are prepared as a suspension in a sterile aqueous diluent, 
determined by the experimental question. Modified Bushnell-Haas broth 
is used as diluent if assaying nutrient- optimized mineralization potential 
(to assess acclimation of the microbial population to the target substrate). 
Ringer's solution is used as diluent if assaying the mineralization potentials 



194 J.E. Lindstrom, J.R Braddock 

of field-treated soils (e.g., fertilized versus unfertilized). Ringer's solution 
may also be amended with macronutrients (N, P), vitamins, or anaerobic 
terminal electron acceptors for bench-scale treatability studies. 

1. Prepare a nominal 1:10 dilution (w/v) of soil in diluent based on soil wet 
mass, preparing a volume sufficient for distribution into several assay 
vials. For example, if three or four replicates are desired per sample, add 
5 g soil to 45 mL diluent. Collect a portion of the soil sample for a dry 
mass determination. The final measured potential will be adjusted per 
gram dry mass accordingly. 

2. Distribute 10 mL of the soil suspension into VOA vials. Prepare a min- 
imum of three replicates for each substrate/treatment combination to 
obtain a mean value for the sample's mineralization potential. Securely 
replace the caps on the vials to avoid gas leakage during incubation and 
C0 2 recovery. 

3. Prepare killed controls ("time zero" samples) to be used for subtracting 
background radioactivity counts from assay samples. Inject 1 mL ION 
NaOH solution through the septum of each control vial. This should 
result in a solution pH above 12 in the vial, halting microbial activity. 
At least three controls should be prepared for each substrate/treatment 
combination, and the mean value is used to "correct" the final result, as 
described below. 

4. Accurately inject 50p.L radiolabeled substrate solution though the sep- 
tum of each vial. Careful measurement is required at this step to assure 
reproducibility of the assay. The injection results in addition of 100 pg 
substrate to the soil suspension. Briefly swirl or shake the vial to mix, 
and incubate under conditions dictated by the experimental design. 

5. Sample microbial activity is terminated at the end of the incubation pe- 
riod (determined from time-course experiments, described below) by 
injecting 1 mL ION NaOH through the septum of each sample vial, as 
described for the time zero controls. Swirl the sample vial to distribute 
the NaOH. Samples may be stored after treatment with NaOH; the high 
pH conditions in the vial sequester the carbon dioxide generated by mi- 
crobial mineralization as carbonates in aqueous solution, preventing loss 
of C0 2 from the vial pending processing to recover the 14 C-labeled C0 2 . 
The samples can be stored for at least a month after this step if necessary. 

Recovery of Evolved 14 C0 2 

Radiolabeled C0 2 evolved from the soil suspension during the incubation 
period is captured by stripping it from solution and capturing it in a basic 
medium. PEA is used to trap the C0 2 . 



9 Assessment of Hydrocarbon Biodegradation 195 

1. Following the incubation period, the soil suspension is acidified by 
adding HC1 to release the C0 2 previously sequestered in solution by 
addition of NaOH. Inject 1.5 mL 12 N HC1 through the septum into the 
VOA vial, and swirl briefly to distribute into solution. 

2. Using the apparatus shown in Fig. 9.1, place the two-hole rubber stopper 
with glass tubing and Tygon on a scintillation vial containing 1 mL PEA 
scintillation cocktail. 

3. Place the influent tubing attached to the scintillation vial on the effluent 
side of the Harvey trap containing acidified toluene. 

4. Attach Tygon tubing to the influent side of the Harvey trap, and attach 
a needle to the other end of the Tygon. 

5. Pierce the septum of the VOA vial with the needle, making certain the 
tip of the needle is above the liquid level in the VOA vial. 

6. Pierce the VOA vial septum with the spinal needle attached to a source 
of N 2 gas. There should be no gas flow until all connections have been 
checked for tightness. 

7. Turn on the N 2 gas source and adjust the gas flow rate to approx. 
lOmL/min. 

8. Strip the C0 2 from the soil suspension for 15min, then stop the gas 
flow through the apparatus. 

9. Remove the stopper from the scintillation vial, place a cap on the vial, 
and determine the amount of radioactivity using a liquid scintillation 
counter. 

1 0. Rinse all glass tubing tips that contacted scintillation cocktail by dipping 
in distilled water several times and wiping clean with a lab wipe; follow 
with an acetone rinse. 

1 1 . Periodically check for radioactivity carryover between samples by run- 
ning method blanks (distilled water in VOA vials) treated as though 
they were samples, except without addition of NaOH or HC1. 

12. After running each sample, check for clogged needles. Spinal needles 
can be cleared with a fine-gauge wire. 

Volatile Versus Nonvolatile Substrates 

If assaying the mineralization potential of a volatile substrate (e.g., benzene, 
toluene, etc.), it is necessary to remove unmetabolized substrate from the 
suspension prior to recovering the C0 2 . This is accomplished by bubbling 
N 2 gas through the suspension after adding the NaOH, but before adding 



196 J.E. Lindstrom, J.R Braddock 

the HC1. The C0 2 will still be sequestered in solution in carbonate form, and 
will not be lost while volatilizing the substrate from the suspension. After 
removing the volatile substrate from the suspension, the C0 2 -stripping 
apparatus is assembled, the sample is acidified, and C0 2 recovery proceeds 
as described. 

Determining Incubation Period 

Relatively high concentrations of both labeled and non-labeled substrate 
are added to the soil suspensions in this assay to avoid interferences from 
field-derived hydrocarbon substrates (Brown et al. 1991). It is therefore 
necessary to detect significant substrate mineralization in a reasonable 
time frame, while avoiding artifacts associated with in vitro acclimation 
of the microbial community assayed. This is accomplished by conducting 
time-course assays with samples prepared as described above. A minimum 
of three replicate assays should be conducted for each incubation period. 

Depending on the substrate chosen, sample incubation times should be 
distributed evenly from time zero to the longest reasonable incubation time. 
Relatively labile substrates (e.g., linear alkanes up to Ci6, low molecular 
weight aromatics up to naphthalenes) may be incubated up to 2 weeks, 
with incubations of, e.g., 0, 3, 7, 10, and 14 days. Anaerobic incubations, 
more recalcitrant substrates, colder temperatures, etc., may dictate time 
courses of longer duration. 

Following completion of the time series, plot and inspect the data. Choose 
an incubation time longer than the observed lag period, but the shortest 
possible time that yields 14 C0 2 recoveries significantly above background 
(time zero data). 

■ Calculation 

The radioactivity recovered from each vial is normalized to a dry soil 
basis, using the data from the portion of soil sample collected for dry mass 
determination. The mean value of the radioactivity recovered from the time 
zero control samples prepared at the beginning of the incubation period 
is then determined, and subtracted from the associated treatment samples 
to obtain a "corrected" radioactivity value for each vial. Note that 1 g wet 
mass of soil is added per vial; thus, each vial represents 1 g wet mass of soil. 

v ^(sample) — ^-(time zero controls) , v 

^(corrected) — TT - ; v^»W 

soil dry mass 

X( corrected) sample radioactivity corrected (Bq/g soil dry mass) 

^(sample) sample radioactivity recovered as C0 2 (Bq) 



9 Assessment of Hydrocarbon Biodegradation 197 

■X(time zero controls) mean radioactivity of controls recovered as C0 2 (Bq) 
Soil dry mass (g dry soil/g wet soil) 

A mean value of radioactivity recovered as 14 C0 2 for each sample can 
be calculated from the corrected Bq/g dry soil data. The radioactivity 
recovered as 14 C0 2 is then compared to that supplied with the added 
labeled substrate. The results maybe expressed as a percentage of substrate 
added that was mineralized in the assay by the formula: 

^ = ^(corrected) ^ m (93) 

^(substrate) 

Si substrate mineralized (%/g soil dry mass) 

X( corrected) radioactivity corrected (Bq/g soil dry mass) 
X( substrate) total radioactivity added to microcosms (Bq) 

Alternatively, the data maybe converted to |ig substrate mineralized. The 
addition of 50 p.L of the substrate solution (2 g/L) results in 100 pg substrate 
being added to the microcosms. The mass of substrate mineralized per 
gram dry soil may be calculated by the formula: 

So x Si , 

S 2 = 9.4 

100 

S 2 substrate mineralized (|ig/g soil dry mass) 
S initial substrate concentration (100 p.g) 

Si substrate mineralized (%/g soil dry mass) 

Depending on the experimental question, results among field treatments 
maybe assessed for significant treatment effects (using unamended diluent 
in the assay). Nutrient-amended assays can be used to demonstrate the 
prior acclimation of microbial communities to hydrocarbon degradation, 
as nutrient limitations potentially present in the field are removed in the 
laboratory assay. Alternatively, a comparison between nutrient-amended 
assays and unamended assays, or among various amendments, may be 
conducted as a treatability study prior to implementing field treatment. 

■ Notes and Points to Watch 

• To assure no gas leakage occurs from the stripping apparatus, all Tygon 
tubing connections (i.e., to glass tubing, Harvey trap, 16-gauge and spinal 
needles) should be secured with several wraps of wire. Tygon tubing can 
be protected from being cut by the wire by wrapping the tubing with 
a piece of laboratory tape before securing with wire. 



198 J.E. Lindstrom, J.R Braddock 

• As noted above, it is important to periodically check for obstructions in 
the various needles and glass tubing used in the stripping apparatus, and 
to clean the glass tubing that comes in contact with scintillation cocktail 
to prevent carryover of radioactivity from previously stripped samples. 

• Carryover of radioactivity from previous samples run on the stripping 
apparatus should be checked periodically by running distilled water 
method blanks. If excessive radioactivity (i.e., significantly above back- 
ground) is recovered from the method blank, change the toluene in 
the Harvey trap, and rerun a method blank. If excessive radioactivity 
persists, it may be necessary to change the Tygon tubing. 

• When using volatile substrates in the assay, the volatile compounds 
require removal prior to recovering the 14 C0 2 , as described. As the 
volatile substrate is radioactive, the exhaust gas from this process must 
be properly captured (e.g., activated carbon filter) and disposed. 

• It is not uncommon to observe substantial variance among samples using 
this assay; careful adherence to the protocol will reduce the variance sub- 
stantially. We recommend preparing as many replicate assays as possible 
in order to obtain a lower standard error for the mean mineralization 
potentials determined. 



References 

Atlas RM (1993) Handbook of microbiological media. CRC Press, Boca Raton, FL 
Bauer JE, Capone DG (1988) Effects of co-occurring aromatic hydrocarbons on degrada- 
tion of individual polycyclic aromatic hydrocarbons in marine sediment slurries. Appl 

Environ Microbiol 54:1649-1655 
B0rresen M, Breedveld GB, Rike AG (2003) Assessment of the biodegradation potential of 

hydrocarbons in contaminated soil from a permafrost site. Cold Regions Sci Technol 

37:137-149 
Braddock JF, Lindstrom JE, Prince RC (2003) Weathering of a subarctic oil spill over 25 years: 

the Caribou-Poker Creeks Research Watershed experiment. Cold Regions Sci Technol 

36:11-23 
Braddock JF, Lindstrom JE, Yeager TR, Rasley BT, Brown EJ (1996) Patterns of microbial 

activity in oiled and unoiled sediments in Prince William Sound. Proceedings of the 

Exxon Valdez Oil Spill Symposium, Feb. 1993. Am Fish Soc Symp 18:94-108 
Brown EJ, Resnick SM, Rebstock C, Luong HV, Lindstrom J (1991) UAF radiorespirometric 

protocol for assessing hydrocarbon mineralization potential in environmental samples. 

Biodegradation 2:121-127 
Collins CH, Lyne PM, Grange JM (1989) Collins and Lyne's microbiological methods, 6 th edn, 

Butterworths, London 
Leahy JG, Colwell RR (1990) Microbial degradation of hydrocarbons in the environment. 

Microbiol Rev 54:305-315 
Lindstrom JE, Prince RC, Clark JC, Grossman MJ, Yeager TR, Braddock JF, Brown EJ (1991) 

Microbial populations and hydrocarbon biodegradation potentials in fertilized shore- 



9 Assessment of Hydrocarbon Biodegradation 199 

line sediments affected by the T/V Exxon Valdez oil spill. Appl Environ Microbiol 
57:2514-2522 

National Research Council (1985) Oil in the sea: inputs, fates, and effects. National Academy 
Press, Washington, DC 

Sugai SF, Schimel JP (1993) Decomposition and biomass incorporation of 14 C-labeled glu- 
cose and phenolics in taiga forest floor: effect of substrate quality, successional state, 
and season. Soil Biol Biochem 25:1379-1389 

Wackett LP, Hershberger CD (2001) Biocatalysis and biodegradation: microbial transfor- 
mation of organic compounds. ASM Press, Washington, DC 

Walker JD, Colwell RR (1976) Measuring the potential activity of hydrocarbon-degrading 
bacteria. Appl Environ Microbiol 31:189-197 



10 



Molecular Techniques for Monitoring 
and Assessing Soil Bioremediation 

Lyle G. Whyte, Charles W. Greer 



10.1 

General Introduction 

Classical culture-dependent microbiological methods have succeeded in 
culturing ~1% of the microbial species in a given environmental sample. 
In reality, this is due to the fact that most isolation procedures are too gen- 
eral, and a wider variety of methods must be developed to recover a larger 
representation of microorganisms from most natural environments. Never- 
theless, our knowledge of microorganisms is largely based on the represen- 
tatives that have been cultured in the laboratory and studied in vitro. Since 
approx. 1990, significant advances in molecular biology techniques have 
transformed environmental microbiology and microbial ecology. These 
techniques bypass the major limitations of culture-dependent microbio- 
logical methods by extracting nucleic acids directly (DNA and RNA) from 
terrestrial or aquatic samples (soils, waters, wastewaters, etc.) and which 
theoretically represent 100% of the microbial species in a given sample. 
A variety of techniques are then used to manipulate and subsequently 
characterize individual DNA and RNA molecules from complex microbial 
communities with a relatively high degree of sensitivity and specificity. 
These techniques have been applied to contaminated soil and aquatic sys- 
tems and have greatly aided in characterizing and monitoring pollutant 
biodegrading microbial populations within these systems. In addition, the 
knowledge gained from using these molecular techniques has helped iden- 
tify novel biodegradation pathways and opened up new perspectives in 
bioremediation processes and pollution abatement. The following survey 
presents an overview of the prominent molecular techniques that are cur- 
rently being utilized for environmental microbiology with a specific focus 
on soil microbiology. The overview is summarized in Fig. 10.1. Several 
specific techniques that include total DNA extraction, polymerase chain 



Lyle G. Whyte: Dept. of Natural Resource Sciences, McGill University, Macdonald Cam- 
pus 21, 111 Lakeshore Road, St. Anne de Bellevue, Quebec, Canada H9X 3V9, E-mail: 
whyte@nrs.mcgill.ca 

Charles W. Greer: Biotechnology Research Institute, National Research Council of Canada, 
6100 Royalmount Ave., Montreal, Quebec, Canada H4P 2R2 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



202 L.G. Whyte, C.W. Greer 

reaction (PCR) analyses, and community characterization using denatur- 
ing gradient gel electrophoresis, are covered in detail in this chapter. 



10.2 

Extraction and Purification of Nucleic Acids (DNA) from Soil 

■ Introduction 

Objectives. All of the current molecular methods crucially rely on the suc- 
cessful extraction and purification of sufficient amounts of nucleic acids 
from environmental samples. Consequently, many methodologies have 
been and continue to be developed for extracting nucleic acids from soils 
and sediments, and improvements are constantly being reported in the 
literature. The soil-DNA-isolation methodologies vary considerably with 
respect to reliability, yield, purity, and degree of shearing, and only recently 
have some of these methods been compared (Yeates et al. 1998; Martin- 
Laurent et al. 2001; Schneegurt et al. 2003; Kauffmann et al. 2004; Mumy and 
Findlay 2004). Several commercial extraction kits are also available such as 
MoBio Laboratories Ultraclean Soil DNA Kit (Mobio Laboratories, Solona 
Beach, CA, USA) and the BiolOl FastDNA soil kit (La Jolla, CA, USA); they 
feature short extraction times and the potential for reduced variability 
(Mumy and Findlay 2004) and have become quite popular despite their rel- 
atively high cost and limitation to the extraction of nucleic acids from 1 g or 
less of sample. In addition, extraction efficiency and resulting nucleic acid 
quality are strongly influenced by the source of the sample, and there are 
numerous co-extracted interfering substances (humics, pollutants, heavy 
metals, etc.). The method described below is routinely used in our lab- 
oratories for isolating nucleic acids from soil and sediments from both 
contaminated and pristine environments, and has yielded fairly uniform 
quantities and qualities of nucleic acids. 

Principle. An initial soil washing step prior to DNA extraction helps solu- 
bilize and reduce contaminants when high quality DNA is required (Fortin 
et al. 2004). Total community DNA is isolated from soil and sediments us- 
ing a direct DNA extraction procedure based on chemical/enzymatic lysis 
(lysozyme and proteinase K in combination with SDS). Released micro- 
bial DNA is ethanol precipitated and the total community DNA is purified 
by a polyvinylpolypyrrolidone (PVPP) spin column filtration step which 
removes contaminating soil organic compounds such as fulvic and humic 
acids (Fortin et al. 2004). 

Theory. The starting point in the analysis of total community DNA from 
environmental samples is the efficient extraction of nucleic acids of suf- 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 



203 



Soil Sample 



Extract Nucleic Acids 
(total community DNA) 



PCR Dependent Analyses 



PCR Amplification 



16S rRNA PCR Fragments 

- Identify, characterize, 
monitor complex 
microbial populations 



Clone Library 

Sequence 

Identification 
of phylotypes 



Catabolic Genotypes 

- Detect, monitor, 
quantitate degradative 
genotypes in 
contaminated soils 



ARDA 

DGGE 

TGGE 

RISA 

RAPD 

- Sequence 
major bands 



PCR Independent Analyses 



Metagenomics 



Metagenomic 
Libraries 

Clone and sequence 
large fragments (BAC 
Fosmids, cosmids) 



Environmental 
Genome Shotgun 
Cloning 

Clone and sequence 
small fragments 



Environmental Microarrays 

Phylogenetic, functional gene 
microarrays to identify, 
characterize, monitor complex 
microbial populations 



Fig. 10.1. Molecular techniques used in environmental microbiology 



ficient quality for subsequent molecular analyses. Two general method- 
ologies are commonly employed: (1) direct lysis of microbial cells within 
the environmental sample by chemical treatments, sonication, freeze-thaw, 
or bead beating protocols; or (2) extraction of the bacterial cells from 
the environmental sample followed by cell lysis. Humic and fulvic acids 
are often co-extracted from soil with the nucleic acids; they must be re- 
moved as they can seriously interfere with subsequent molecular reactions 
(DNA polymerase amplification, DNA-DNA hybridizations, DNA labeling, 
restriction nuclease digestion). The released soil DNA usually can be puri- 



204 L.G. Whyte, C.W. Greer 

fied by a variety of methods/techniques such as chromatography and silica 
gel or PVPP spin-filter columns. The quality and quantity of the purified 
soil DNA extract is generally verified by agarose gel electrophoresis and/or 
spectrophotometry (Abs. 260/280 nm). 

■ Equipment 

Water bath incubators (30 °C, 37 °C, 85 °C) 

Microcentrifuge (13,600-15,800 g) at4°C 

Platform shaker 

Speedvacuum (optional) 

Gel electrophoresis apparatus, ultraviolet light source, and camera or gel 
documentation system 

Spectrophotometer, quartz cuvettes 

Vortex 

Pipettors (10-20, 100, 1,000 pL) and tips 

Microcentrifuge tubes (0.5, 1.5, 2.0 mL) 

MicroSpin columns (Amersham Biosciences, Baie d'Urfe, Que., Canada) 

Reagents 

Buffer 1: 50 mM Tris-HCl, pH 8.3, containing 200 mM NaCl, 5 mM EDTA, 
and 0.05% Triton X-100 (Fisher Scientific, Nepean, Ont. Canada) 

Buffer 2: 50 mM Tris-HCl, pH 8.3, containing 200 mM NaCl and 5mM 
EDTA 

Buffer 3: 10 mM Tris-HCl, pH 8.3, containing 0.1 mM EDTA 

Distilled water 

250 mM Tris-HCl, pH 8.0, with 5 mg/mL lysozyme (freshly prepared) 

Proteinase K (20 mg/mL) 

20% SDS 

7.5 M ammonium acetate 

2-propanol 

70% ethanol 

TE buffer: 10 mM Tris-HCl, 1 mM EDTA, pH 8.0 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 205 

• TAE buffer: 40 mM Tris-acetate, 1 mM EDTA, pH 8.0 

• Acid-washed PVPP spin columns (preparation below) 

• 200 mM potassium phosphate buffer, pH 7.0 

• Cone. HC1 

Note: All solutions should be sterilized by filtration or autoclave. The recipes 
for many of the solutions described throughout this chapter can be found 
in Sambrook and Russell (2001). 



■ Sample Preparation 

Soil samples to be extracted should be frozen (-20 or -80 °C) as soon as 
possible upon collection and stored frozen to minimize changes in micro- 
bial communities because of the sampling process and/or degradation of 
nucleic acids. 



■ Procedure 

Soil-Washing Step 

1. Add 1 mL of buffer 1 to 0.5 g of soil or sediment in a microcentrifuge 
tube. Mix by vortexing (speed 4) for 30 s, then by inverting for 1.5 min. 

2. Centrifuge 5 min at 4 °C at 3,000 g. Remove the supernatant with a pipette. 

3. Add 1 mL of buffer 2 to sediment. Mix by vortexing (speed 4) for 30 s, 
then by inverting for 1.5 min. 

4. Centrifuge 5 min at 4 °C at 3,000 g. Remove the supernatant with a pipette. 

5. Add 1 mL of buffer 3 to sediment. Mix by vortexing (speed 4) for 30 s, 
then by inverting for 1.5 min. 

6. Centrifuge 5 min at 4 ° C at 3,000 g. Remove the supernatant with a pipette. 

Nucleic Acid Extraction Step 

1. Add 450 p.L of sterilized distilled water to the 0.5 g of sample in a 1.5 mL 
microcentrifuge tube. Vortex (moderate speed) for approx. 2 s to dis- 
lodge the pellet. 

2. Add 50 pL of 250 mM Tris-HCl, pH 8.0, containing lysozyme (5 mg/mL). 

3. Incubate at 30 °C, mixing by inversion, for 30 min. 

4. Transfer to a 37 °C water bath and incubate for another 30 min, mixing 
by inversion every 10 min. 



206 L.G. Whyte, C.W. Greer 

5. Add 5 pL of proteinase K (20 mg/mL). Incubate for 1 h at 37 °C, mixing 
by inversion every 10 min. 

6. Add 50 pi of 20% SDS. Incubate at 85 °C for 30 min, mixing gently by 
inversion every 10 min. 

7. Centrifuge 10 min at room temperature at 13,600 g. Transfer the super- 
natant to a fresh 1.5 mL microcentrifuge tube. 

8. Add 1/2 volume of 7.5 M ammonium acetate. Mix gently by inversion, 
and incubate on ice for 15 min. 

9. Centrifuge 5 min at 4 °C, at 13,600 g. Transfer and split the supernatant 
in two fresh 1.5 mL microcentrifuge tubes and treat each tube sepa- 
rately. 

10. Add 1 volume of cold 2-propanol, and precipitate DNA overnight at 
-20 °C. 

11. Centrifuge 15 min at 4 °C at 15,800 g. Discard the supernatant. 

12. Wash the pellet with 500 pL of cold 70% ethanol. Gently tap the tube or 
mix by inversion. 

13. Centrifuge 5 min at 4 °C at 15,800 g. Discard the supernatant. 

14. Dry the pellet in speedvacuum for 5 min or air dry the pellet (approx. 
30 min). 

15. Add 100 pi of TE buffer to the pellet in each tube, place the sample 
on ice on a shaking platform and let the pellet slowly dissolve (approx. 
1 h). Combine the DNA extracts from the two tubes. 

16. Warm up the DNA combined extract for 10 min at 37 °C. 

17. Purify 50 pL of total community DNA on a PVPP column. 



DNA Purification with PVPP Columns (Modified from Berthelet et al. 1996) 

1. Prepare acid washed PVPP. Pour 1,034 mL of cone. HC1 (11.6M) slowly 
with stirring into 2,966 mL of MilliQ (Qiagen Inc., Hilden, Germany) 
water; this will result in ca. 4 L of 3 M HC1. Add 150 g PVPP and suspend 
with stirring at room temperature for 12-16 h. Leave the suspension 
to settle for 30-60 min, then aspirate or decant the supernatant. Again 
suspend the PVPP, now in approx. 3.5 L of 200 mM potassium phosphate 
buffer (pH 7.0) and stir 1-2 h. Repeat the aspiration/decant and suspen- 
sion twice more until the supernatant pH is close to 7.0 (check aliquot 
with pH meter). Then repeat the aspiration/decant and suspension two 
more times with approx. 3.5 L of 20 mM potassium phosphate buffer 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 207 

(pH 7.0). Aliquot the final suspension into small bottles and autoclave 
(15-20 min, 121 °C). Store at 4°C. 

2. Acid washed PVPP (0.9 mL) slurried in 20 mM potassium phosphate 
(pH 7.0) is added to empty sterile MicroSpin columns placed inside 2.0- 
mL tubes and centrifuged for 3 min at 735 g at room temperature. If the 
top of the column is still immersed following centrifugation, remove the 
liquid from the collection tube, and re-spin the column. 

3. Load the "crude" DNA extract (50-100 p.L) onto the center of the column 
being careful not to touch the side of the column. This ensures that all 
of the sample will pass through the column and be cleaned, and not run 
down the side of the column. 

4. Place the loaded columns in the microcentrifuge, ensuring that the slop- 
ing face of the packed column is facing the middle of the centrifuge. 
Centrifuge the columns for 3 min at 735 g at room temperature and 
collect the filtrate. 

5. The "clean" DNA extract is then stored at -20 °C and is ready for PCR. 
The used PVPP is discarded and the MicroSpin columns washed for 
reuse. 



Agarose Gel Electrophoresis 

We check the quality and quantity of ca. 5 pL of purified soil DNA extract 
by both agarose gel electrophoresis in 0.7% gels in TAE buffer (stained with 
ethidium bromide and visualized by ultraviolet light) and spectrophotom- 
etry (Abs. 260/280 nm) using standard methods as described by Sambrook 
and Russell (2001). 



Notes and Points to Watch 

This methodology can be readily scaled up to 10 g soil samples as de- 
scribed in Fortin et al. (2004). 

All molecular biology methodologies are notoriously variable. Incuba- 
tion temperatures and durations, volumes, centrifugation parameters, 
etc. should be vigorously adhered to. 

Lysis of the microbial cells during DNA extraction represents a critical 
step in PCR-mediated approaches (von Wintzingerode et al. 1997). Each 
physical, chemical, and biological step involved in the preparation and 
analysis of an environmental sample is a source of bias which might give 
a distorted view of a given ecological niche (von Wintzingerode et al. 
1997). It is often a question of whether there was sufficient or preferential 



208 L.G. Whyte, C.W. Greer 

disruption of microbial cells. Rigorous conditions maybe required to lyse 
Gram-positive cells but also may cause excessive shearing of nucleic acids 
of the Gram-negative cells, potentially biasing the reported diversity 
of the sample as well as possibly creating artifacts and chimeric PCR 
products (Liesack et al. 1991). Therefore, checking the soil DNA extract 
by agarose gel electrophoresis will indicate the quality of the extracted 
DNA and the extent of shearing. 

• Quantification of the soil DNA extract by spectrophotometry is often 
inaccurate and does not appear to correlate with other methods such as 
agarose gel electrophoresis using known concentrations of DNA stan- 
dards or PicoGreen (Molecular Probes, Leiden, Netherlands). 



10.3 

Amplification of Catabolic Genotypes 

and 16S rDNA Genotypes by PCR 

■ Introduction 

Objectives. Many of the molecular methodologies used in environmental 
microbiology rely on a PCR amplification step and are therefore considered 
PCR dependent. The objective of PCR is to amplify target gene sequences 
from total community DNA extracted from an environmental sample. The 
amplified sequences can then be characterized by a variety of molecular 
methodologies as shown in Fig. 10.1. In soil biodegradation studies, the 
target gene sequences can be either catabolic (biodegradative) genes of in- 
terest or a phylogenetic gene (almost always the 16S rDNA gene). In the case 
of catabolic genes, the PCR amplification step is generally used to detect 
the presence or absence of various catabolic genotypes in contaminated 
soils. Determining the prevalence and composition of specific biodegrada- 
tive genotypes, and hence microbial populations, in contaminated soils 
significantly aids in assessing the feasibility of using biotreatment and in 
developing appropriate bioremediation strategies for a particular contam- 
inated site, as well as in monitoring the effects on specific populations 
during bioremediation operations. For example, we routinely use PCR 
screening for hydrocarbon-degradative genotypes to perform biotreatabil- 
ity assessments of contaminated soils (Whyte et al. 1999; Soloway et al. 200 1 ; 
Whyte et al. 2001) and to monitor bioremediation treatments (Whyte et al. 
2003). The prevalence of various alkane monooxygenase genotypes and 
other degradative genotypes in hydrocarbon-contaminated and pristine 
soils from a variety of Arctic, Antarctic, and alpine contaminated soils was 
also determined by PCR screening (Whyte et al. 2002; Margesin et al. 2003; 
Luz et al. 2004). 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 209 

In comparison, PCR amplification is currently the most widely used 
method to obtain 16S rDNA genotypes for detailed characterization of 
microbial communities. Unlike PCR amplification of degradative geno- 
types, however, PCR amplified 16S rDNA genes from a total commu- 
nity DNA extract must be further characterized by one or more of the 
molecular methodologies [clone and sequencing, denaturing gradient gel 
electrophoresis (DGGE; see Sect. 10.4, below), temperature gradient gel 
electrophoresis (TGGE), etc.], shown in Fig. 10.1, to obtain meaningful 
information on the amplified 16S rDNA PCR product. These molecular 
methods have been used to characterize cold-adapted microbial popula- 
tions in hydrocarbon-contaminated soils originating in northern Canada 
( Juck et al. 2000), to determine the effect of oil contamination and a biostim- 
ulation treatment on Pseudomonas diversity in soil microcosms (Evans et 
al. 2004), to monitor microbial population changes in beach sediments dur- 
ing an experimental oil spill (Macnaughton et al. 1999), and to monitor the 
impact on microbial community composition during the bioremediation 
of hydrocarbon-contaminated soils (Mills et al. 2003). 

Principle. During PCR, double-stranded DNA (from total community DNA 
soil extracts) is separated into single strands at high temperature (denat- 
uration). Two oligonucleotide primers then anneal (at a lower annealing 
temperature) to complementary regions (which flank the target sequence) 
of the single-stranded DNA. A heat-stable DNA polymerase synthesizes 
a new strand of DNA by extending the primer using the complementary 
strand as a template, thus creating a duplicate copy of the target sequence. 
This cycle is repeated 20-30 times resulting in an exponential amplification 
(2 20 -2 30 fold) of the target sequence. 

Theory. PCR is the simplest and currently the most widely used method 
to detect/obtain catabolic genotypes or 16S rDNA genotypes for detailed 
downstream characterization of soil microbial communities. These proce- 
dures are increasingly being utilized to perform biotreatability assessments 
of contaminated soils, to monitor the effects of soil bioremediation treat- 
ments on microbial populations, and to identify and characterize important 
and/or novel biodegradative microbial strains or groups of microorganisms 
in contaminated soils. This Section describes PCR procedures for the am- 
plification of catabolic genotypes and 16S rDNA genes for cloning and 
sequencing. 



Equipment 

PCR work station chamber (UV hood; optional but recommended for 
16S rDNA PCR) 



210 L.G. Whyte, C.W. Greer 

• Microcentrifuge 

• Thin walled 0.2-mL PCR tubes 

• Pipettes (10, 100 pL) and tips (filtered tips are recommended for 16S 
rDNA PCR) 

• PCR thermocycler 

• Gel electrophoresis apparatus, ultraviolet light source, camera or gel 
documentation system 

• PCR cleanup kit (QIAquick PCR Purification Kit; Qiagen Inc.) 

• PCR cloning kit (Promega pGem-T Easy Cloning Kit; Promega Corpo- 
ration, Madison, WI, USA) 

• Petri dishes 

• Incubator (37 °C) 

• Water bath incubators (37, 42 °C) 

■ Reagents 

Catabolic Genotype PCR Amplification 

• Catabolic target gene forward and reverse oligonucleotide primers (for- 
ward and reverse; 0.4-0.8 mM stock solutions) 

• Soil DNA extract 

• DNA from reference organisms 

• 100 bp DNA ladder (Fermentas, SM0241, Invitrogen, Carlsbad, CA, USA) 

• DNA polymerase (Taq DNA polymerase is often used) 

• DNA polymerase buffer: 10 mM Tris-HCl, pH 9.0, containing 50 mM KC1 
and 15mMMgCl 2 

• 25 mM MgCl 2 

• 1.25 mM stock solution of each deoxynucleoside triphosphate (dNTP), 
namely dATP, dCTP, dGTP, dTTP 

16S rDNA PCR Amplification 

• General ("universal") Bacteria primers: 

- 27F(10pM)5 / -GGTTACCTTGTTACGACTT 

- 758R (10 pM) y-CTACCAGGGTATCTAATCC 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 211 

Soil DNA extract 

DNA from reference organisms (control DNA) 

100 bp DNA ladder (Fermentas, SM0241) 

Taq DNA polymerase (5 U/pL; Invitrogen) 

10 x PCR buffer: 200 mM Tris-HCl/500 mM KC1 

50 mM MgCl 2 

10 mM stock solution of each dNTP (dATP, dCTP, dGTP, dTTP) 

Bovine serum albumin (BSA; 10 mg/mL in sterile UV irradiated ddH 2 0) 

Sterile UV irradiated H 2 

Store all PCR reagents at -20 °C. 

Cloning and Sequencing of 16S rDNA PCR Amplicons: Transformation 

• Lac~ competent E. coli cells (DH5a) 

• Isopropyl-B-D-thiogalactopyranoside (IPTG) solution: 0.1 M, filter ster- 
ilized (store at 4 °C) 

• Ampicillin (Amp) solution: 10 mg/mL, filter sterilized (store at 4°C) 

• 5-bromo-4-chloro-3-indoyl-/J-d-galactoside (X-Gal) solution: 50 mg/mL 
in N, N'-dimethyl-formamide, make fresh for each transformation (store 
at-20°C) 

• Luria broth (LB) medium (per L): 10 g tryptone, 5g yeast extract, 5g 
NaCl, adjust to pH 7.0 with NaOH 

• LB plates with Amp/IPTG/X-Gal: To 1 L of autoclaved LB medium, add 
5 ml of IPTG solution, 10 ml of ampicillin solution, and 1.6 mL of X-Gal 
solution. 

Note: All solutions should be sterilized by filtration or autoclave. The recipes 
for many of the solutions described throughout this chapter can be found 
in Sambrook and Russell (2001). 

■ Sample Preparation 

Total community DNA is extracted and purified as described in Sect. 10.2. 
The soil DNA extracts should always be stored frozen at -20 °C or -80 °C 
and kept on ice during the procedure to minimize nucleic acid degradation, 
which will occur at greater rates at higher temperatures. 



212 L.G. Whyte, C.W. Greer 

■ Procedure 

Primer Design 

An important key to PCR is optimal design of oligonucleotide primers 
specific to the desired gene target of interest. It is the specificity of the 
primers that allows PCR to amplify catabolic or 16S rDNA genes that can 
be in low abundance in complex environmental samples. 

Catabolic Gene PCR Primers 

We have designed and utilized a variety of primers for PCR amplification 
of catabolic genes; the specific oligonucleotide sequences of these primers 
are available for a variety of hydrocarbon degradative genes (Whyte et al. 
2002; Margesin et al. 2003; Luz et al. 2004) and dehalogenation of chlori- 
nated organics (Fortin et al. 1998). In general, we design PCR primers as 
follows. Gene sequences for key enzymes from known bacterial biodegrada- 
tive pathways are identified and searched for in databases such as the nu- 
cleotide database (GenBank) at the NCBI web site (www.ncbi.nlm.nih.gov). 
The DNA sequences of all corresponding genes encoding the key enzyme 
are retrieved for comparative DNA and protein alignments using appropri- 
ate molecular biology sequence software. PCR forward and reverse primers 
for each catabolic gene sequence are then selected from the alignments by 
PCR primer software and/or manually by identifying homologous regions 
shared by the selected DNA sequences. For construction of the oligonu- 
cleotide primers, long sequences originating from within the coding region 
of the catabolic genes and having a high G+C content are preferred to ensure 
specificity. In addition, we use the following general criteria for designing 
catabolic gene primers: 

- Generally 20-30 nt in length. 

- At least 5 nt at both ends of the primer exhibiting exact match pairing 
with the target DNA sequence, 

- Ideally, if there are mismatches, they should be in the middle of the 
primer sequence. 

- Ideally, the ends of the primer should terminate with 2-3 G or C. 

- Both forward and reverse primers should possess similar G+C content, 
with an average G+C content of 40-60%, with limited stretches of poly- 
purines or polypyrimidines to ensure specificity. 

- The PCR products generated should be ca. 200- 1,000 nt in length and 
thus produce an easily detectable band by agarose gel electrophoresis. 

Specificity of the selected primer sequences for the gene of interest is 
then verified by Fasta and blastn search programs available at the NCBI 
website. 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 213 

16S rDNA PCR Primers 

Given the varying degrees of conservation of thel6S rDNA gene for Bacteria 
and Archaea, 16S rDNA primers or probes can be designed with any degree 
of specificity for groups, ranging from both domains (Ward et al. 1992), 
to a single domain (Battin et al. 2001), or to various subgroups all the way 
down to the species and sub-species level (Ahn et al. 2002). Generally, most 
16S rDNA PCR-based studies rely on using a set of general or "universal" 
primers specific for the Bacterial domain and/or less often, the Archaeal 
domain. These general primer sets are readily available in the literature but 
should be updated periodically as the 16S rDNA database grows daily. One 
can also design new conserved 16S rDNA primer sets by accessing the Ribo- 
somal Database Project (RDP) website (http://rdp.cme.msu.edu/index.jsp) 
algorithm that can be used in choosing the proper primer set for the PCR 
amplification in question. 



PCR Amplification, Cloning, and Sequencing 
PCR Amplification of Catabolic Genotypes 

1. For each PCR reaction, set up the following reaction in a 0.2-mL micro- 
centrifuge tube: 

- 1-5 pi of total community DNA soil extract (ca. lOOngofDNA) 

- 2 pL of each oligonucleotide primer (0.4-0.8 mM stock; final concen- 
tration 0.2 pM) 

- 5 pL of DNA polymerase buffer 

- 2pLof25mMMgCl 2 

- 8 pL of 1.25 mM dNTP (200 pM of each dATP, dCTP, dGTP, and dTTP 
solution) 

- 2.0-5.0 U of DNA polymerase 

2. The final volume in the tube is brought to 50 pL with sterile distilled 
water. Prior to the addition of DNA polymerase, the samples are boiled 
for 2 min and then transferred to ice. 

3. Negative control: The same mixture is used for the negative control 
except that the total community DNA soil extract is replaced with sterile 
distilled water. 

Positive Control: The same mixture is used for the positive control except 
that the total community DNA soil extract is replaced with a genomic 
DNA extract from the appropriate control reference organism. Template 
DNA for PCR from the reference organism can be obtained by resus- 
pending 2-3 colonies in 500 pL of sterile distilled water and boiling for 



214 L.G. Whyte, C.W. Greer 

10 min. The sample is cooled on ice, centrifuged in a microcentrifuge for 
2 min at 12,000 g, the supernatant collected, and stored at -20 °C. 

4. PCR is conducted using an appropriate PCR thermal cycler. We generally 
are successful using the following PCR parameters: 

- 30 cycles of 1 min at 94 °C (denature) 

- 1 min at 60 °C (anneal) 

- 1 min at 72 °C (extend) 

- A final extension of 3 min at 72 °C. 

5. To determine the presence or absence of the appropriately sized PCR 
fragment, ca. 5-10 pL of the PCR reaction mixture of soil DNA extracts 
and the corresponding positive and negative controls, and a 100 bp DNA 
ladder are analysed by agarose gel electrophoresis (1-1.4% agarose gels 
using TAE buffer) and visualized by ethidium bromide staining essen- 
tially as described by Sambrook and Russell (2001). There should be 
a single band of the same size in both the positive and sample lanes, with 
no band in the negative control lane. 

6. To confirm that DNA had been successfully extracted from the soils and 
could be amplified by PCR, general ("universal") 16S rDNA bacterial 
primers are used as a positive PCR amplification control for all soil DNA 
extracts (Whyte et al. 2002). 

PCR Amplification of 16S rRNA for Downstream Cloning and Sequencing 

The protocol given is to PCR amplify 16S rDNA from an environmental 
sample. 

1. All subsequent work should be conducted in an enclosed PCR work 
station chamber. This will help eliminate contamination of plastic ware 
with extraneous 16S rDNA that is present ubiquitously. Latex gloves 
should be worn throughout the procedure. 

2. A PCR master mix (enough for three PCR reactions) is prepared in 
a 1.5 mL microcentrifuge tube by combining the following reagents: 

- 7.5 pL of each oligonucleotide primer (10 pM stock; final concentra- 
tion 0.5 pM) 

- lpLoflOx PCR buffer 

- 4.5 pL of 50 mM MgCl 2 (final concentration 1.5 mM) 

- 3pL of 10 mM dNTP stock solution (final concentration 200 pM of 
each dATP, dCTP, dGTP, and dTTP) 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 215 

- 1.9pLofBSA(10mg/mL) 

- 92.6 pL of sterile, irradiated water 

3. Briefly microcentrifuge the PCR master mix tube at 15,000 g for ca. 10 s. 

4. Add 3 pL of Taq polymerase to the PCR master mix tube, briefly vortex 
(2-4 s) to mix and microcentrifuge at 15,000 g for ca. 10 s. 

5. Label the following 0.2 mL PCR reaction tubes: positive control, negative 
control, sample. To each reaction tube add 45 pL of master mix. 

6. Add 5 pL of E. coli (or other positive control) genomic DNA (ca. 50- 
100 ng) to the positive control. To the negative control, add 5 pi of water. 
To the sample tube, add 1-5 pL of total community DNA soil extract (ca. 
100 ng of DNA). 

7. Briefly vortex (2-4 s) the PCR reaction tubes to mix. Microcentrifuge the 
tubes at 15,000 g for ca. 10 s. 

8. PCR is conducted in an appropriate thermal cycler. We are generally 
successful using the following PCR parameters: 

The first 10 cycles are conducted using a "touchdown protocol" from 
65-55 °C, with the annealing temperature decreasing by 1 °C at each 
cycle. 

- lminat94°C 

- lmin at 65-55 °C 

- 3minat72°C 

The subsequent 20 cycles are performed with an annealing tempera- 
ture of 55° C. 

9. The presence or absence of the appropriately sized PCR fragment (73 1 bp 
for Bacteria 16S rDNA) is determined by agarose gel electrophoresis 
(0.8%) of 5 pL of the PCR reaction mixture of soil DNA extracts, posi- 
tive and negative controls, and a 100 bp DNA ladder, and visualized by 
ethidium bromide staining essentially as described by Sambrook and 
Russell (2001). There should be a single band of the same size in both 
the positive and sample lanes, with no band in the negative control lane. 

Cloning and Sequencing of 16S rDNA PCR Products 

1 . We generally use a spin column purification system such as the Qiaquick 
PCR purification kit to clean the PCR reaction prior to cloning. It is 
important to purify PCR reaction products as unbound primers and 
unincorporated nucleotides can be inhibitory to the ligation reaction. 

2. The purified PCR product is quantified by spectrophotometry (Abs. 
260/280 nm; Sambrook and Russell 2001). 



216 L.G. Whyte, C.W. Greer 

3. Because of their ease of use and reliability, commercial PCR cloning 
kits, such as the pGEM-T Easy Vector (Promega Corp.), are commonly 
used for ligating 1 6S rDNA PCR amplicon libraries into a cloning vector. 
A ligation reaction is set up as described in the pGEM-T Easy Vector 
technical manual. If this is a first-time attempt at cloning with this 
PCR product, it may be necessary to optimize the vectoninsert molar 
ratio. We generally optimize the reaction with 1:3, 1:1, and 3:1 ratios. 
To calculate the amount of PCR product to include in the reaction use 
the following formula: 

ng insert 

(50 ng of vector) x (size of PCR product) x (insert:vector ratio) 
kb size of vector (3.0 kb for pGEM-T Easy) 

4. Incubate the ligation reaction at 4 °C overnight. 

5. Transformation protocols of E. coli competent cells with recombinant 
vector (16S rDNA inserted into the pGEM-T Easy Vector) can be found 
in the pGEM-T Easy technical manual or Sambrook and Russell (2001). 
Competent cells can be provided with the pGEM-T Easy Vector system; 
we have had success using E. coli DH5a (made chemically competent 
as described by Sambrook and Russell 2001). We generally follow the 
protocol provided in the manual. Always ensure that positive and neg- 
ative controls are included in the analysis. The positive control consists 
of cells transformed with the vector DNA alone; the negative control 
cells are treated the same as cells being transformed, but with no added 
DNA. 

6. Spread plate 100 p.L of each transformation (in duplicate) and controls 
onto appropriately labeled LB/Amp/X-Gal/IPTG plates. 

7. Incubate plates overnight at 37 °C. 

8. Score blue and white colonies. White colonies arise from insertion of 
a cloned product into the pGEM-T Easy Vector. More than 60% white 
colonies should be observed. 

9. White colonies are either directly sequenced or screened for unique 
clones prior to sequencing by amplified ribosomal DNA restriction 
analysis (ARDRA; sometimes called restriction fragment length poly- 
morphism, RFLP; see Massol-Deya et al. 1997 for a typical ARDRA 
protocol for 16S rDNA amplicons). Sequencing of the 16S rDNA in- 
serts in the pGEM-T Easy Vector system is conducted with primers as 
described by that system's technical manual; ensure that when ream- 
plifying the cloned inserts to use the pGemT-Easy primers T7 and Sp6 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 217 

to avoid amplifying E. coli 16S rDNA genes. Sequencing is usually per- 
formed by commercial laboratories or by in-house sequencing facilities 
commonly found in most large research institutions. 

10. Sequences are submitted for comparison and identification to the Gen- 
Bank databases using the NCBI Blastn algorithm, the EMBL databases 
using the Fasta algorithm (http://www.ebi.ac.uk/fasta33/nucleotide. 
htmL) and/or the Ribosomal Database Project (RDP) using its Sequence 
Match. Sequences that demonstrate strong homology are then aligned 
to reference sequences and phylogenetic trees commonly constructed 
( Juck et al. 2000). Sequences that demonstrate uncertain alignments are 
checked for chimeras using the CHECK_CHIMERA software program 
function at the RDP site. 



Notes and Points to Watch 

A key limitation to 16S rDNA PCR amplifications is contamination of 
DNA introduced by unintentional tube-to-tube contamination or con- 
taminated reagents. For this reason, false-positive signals and false- 
negative amplifications are not uncommon due to the extreme sensitivity 
of the 16S rDNA PCR reaction, and the ubiquity of 16S rDNA genes in 
almost all biological materials. Fortunately, this problem can be avoided 
simply by using good laboratory techniques as indicated above. 

We often perform PCR amplification on both undiluted soil DNA extracts 
(as described here) and diluted extracts (1/10, 1/100). Diluting the DNA 
extract can result in the parallel dilution of undesired contaminants that 
inhibit the PCR reaction; it is not uncommon to observe successful PCR 
amplification from the diluted samples but not the undiluted sample. 

To minimize the loss of nucleic acids from small sample volumes, ad- 
ditives such as BSA and T4 gene 32 (gp32) can be used to reduce the 
inhibitory effect of contaminants (Kreader 1996). 

The PCR procedures described here should be considered qualitative 
rather than quantitative. Differences in band intensity do suggest dif- 
ferences in the relative amounts of the genotypes in the original sam- 
ples, but keep in mind that PCR reactions are very sensitive to reaction 
conditions. Quantitative PCR protocols (real-time quantitative PCR or 
RT-qPCR) have been recently developed and are being applied to con- 
taminated soils. 

Very similar nucleic acid sequences can also affect amplification of to- 
tal community DNA, especially during 16S rDNA PCR amplifications. 
Chimeric sequences result from the heterologous combination of two 



218 L.G. Whyte, C.W. Greer 

non-identical but similar strands of DNA, but do not generally exist 
in the sample being investigated. However, chimeric sequences can be 
formed at frequencies of several percent during PCR (Liesack et al. 1991). 
The resultant PCR artifacts can affect subsequent analyses by erroneously 
suggesting the existence of novel taxa from these hybrid sequences. The 
binding of heterologous DNA into chimeric structures has also been 
shown to compete with the binding of specific primers during the an- 
nealing step (Meyerhans et al. 1990; Ford et al. 1994; Wang and Wang 
1996). As well, DNA damage such as that caused by mechanical and 
chemical shearing has been suggested to contribute to the formation of 
chimeric DNA during PCR (Paabo et al. 1990). 

• Another pitfall of PCR is the production of minute errors by Taq DNA 
polymerase, which lacks the ability to proofread. (Ford et al. 1994). 
However, this is only a potential problem when sequencing the resulting 
PCR products. 

• For cloning into the pGEM-T Easy Vector system, it is essential to use 
a thermostable polymerase that lacks 3 / -5 / exonuclease activity in the 
initial 16S rDNA amplification step of soil DNA extracts. This will in- 
sure that a 3'A overhang is present on the PCR product and will greatly 
improve the efficiency of the ligation process, as well as avoiding cir- 
cularization of PCR products. Common polymerases that lack the 3 f -5 f 
exonuclease activity are Taq, Tfl, and Tth. 



10.4 

DGGE Analysis Soil Microbial Communities 

■ Introduction 

Objectives. Denaturing gradient gel electrophoresis (DGGE) is a very ver- 
satile method for screening the total microbial community DNA from 
a complex sample. Our limited knowledge of the total microbial com- 
munity composition and function in complex environmental samples has 
necessitated the development of techniques like DGGE to enable us to look 
more directly at the representative microorganisms, independent of the bi- 
ases introduced by culturing. In the last 10 years, more than 1,000 articles 
have been published using DGGE for the analysis of various environmental 
samples. 

DGGE analysis of microbial communities produces a complex profile 
or banding pattern, which can be quite sensitive to spatial and temporal 
sampling variations (Murray et al. 1998). The classic means of analyzing this 
variability has been visual, reporting differences between samples in band 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 219 

intensity, or the presence or absence of specific bands. However, a recent 
study suggests that the results of denaturing gradient methods are readily 
amenable to statistical analysis, provided there is sufficient standardization 
of analytical procedures (Fromin et al. 2002). This would provide the rigor 
of statistical validation of observations and permit a broader range of 
comparisons to be made between different samples and between different 
experimental or environmental parameters. 

DGGE is a useful method for visualizing the major members of a micro- 
bial community, but several factors must be considered when interpreting 
the data. The limit of resolution of this method is about 1% of the total 
community population (Muyzer et al. 1993; Murray et al. 1996), and in very 
complex samples, more bands may be produced than can be resolved. Ini- 
tial calibration to ensure optimal gradient and electrophoretic conditions 
is also important (Muyzer et al. 1993; Muyzer and Smalla 1998). DGGE 
requires rather large quantities of DNA for reliable visualization, possibly 
as much as 500 ng for environmental samples (Nakagawa and Fukui 2002). 
Also, DGGE is typically limited to fragments of no more than 500 bp (My- 
ers et al. 1985), which limits the amount of sequence information that can 
ultimately be retrieved. Some ambiguity can exist in associating a single 
band in a DGGE profile with a single microbial species, since it is possible 
that multiple amplicons co-migrate to the same location in the gel, and 
similarly, multiple bands may be produced by a single species since multi- 
ple copies of 16S rDNA do exist in the same microorganism (Nubel et al. 
1997). 

Principle. DGGE separates a mixture of PCR-amplified DNA fragments 
according to differences in sequence G-C content, based on their differential 
mobility through a DNA-denaturing gel. Once separated, the individual 
fragments can be recovered from the gel and the nucleotide sequences 
determined and compared against existing databases (GenBank, Ribosome 
Database Project) to identity microorganisms in the sample. 

Theory. DGGE, which is based on the early work of Fischer and Lerman 
(1979), is one of the most commonly used methods for the characterization 
of complex microbial communities, and was pioneered by Muyzer et al. 
(1993) for environmental samples. In a manner similar to the other PCR- 
based characterization techniques, samples for DGGE analysis are prepared 
either directly from PCR-amplified environmental DNA (Ahn et al. 2002; 
Ibekwe et al. 2002), from clone libraries constructed from PCR-amplified 
environmental samples (Liu et al. 2002), or in some cases from colonies 
obtained from enrichment cultures (Bonin et al. 2002). Total community 
DNA is extracted, purified and used as a PCR template for the amplification 
of specific target molecules. The most common target molecule is the 16S 
rDNA gene which is used as a phylogenetic marker to assess biodiversity 



220 L.G. Whyte, C.W. Greer 

and eventually to identify individual members within the community. Gen- 
eral 16S rDNA primers, often referred to as universal primers, are used to 
amplify the total community DNA. This produces a mixture of fragments 
derived from the individual microorganisms in the sample. Because each 
fragment has a different internal sequence, the fragments can subsequently 
be separated based on their melting behavior in a denaturing gradient, usu- 
ally composed of urea and formamide. As the double-stranded PCR frag- 
ments move through the gel from low to high denaturant concentration, 
they begin to separate into single strands, which reduces their mobility. 
Complete strand separation is prevented by incorporating a GC rich region 
(ca. 40 bases), referred to as a GC clamp, at the 5'-end of one of the PCR 
primers. The DNA comes to rest when it is almost fully denatured. The 
position along the gradient at which the DNA stops is determined primar- 
ily by the relative proportions of G+C and A+T in a given amplicon, since 
G-C bonds are more difficult to denature than A-T bonds. Properly cali- 
brated, DGGE is sensitive enough to detect even single base-pair differences 
between amplicons (Miller et al. 1999). The result in complex samples is 
typically a banding pattern that is representative of the molecular diversity 
in the sample. The individual bands can subsequently be extracted from 
the DGGE gel and sequenced to potentially identify individual microor- 
ganisms. 

■ Equipment 

• See Sect. 10.3 for equipment for PCR amplification 

• Gradient mixer (BioRad Model 385 Gradient Former, BioRad Laborato- 
ries Inc., Mississauga, Ont. Canada) 

• BioRad Dcode Universal Mutation Detection System (BioRad Labs.; or 
equivalent) 

• Fluorlmager system, model 595 (Molecular Dynamics Inc., Sunnyvale, 
CA, USA; or equivalent) 

• PCR clean up kit (QIAquick PCR Purification Kit, Qiagen Inc.; containing 
PB, EB, PE buffer, and column- collection tubes) 



Reagents 

50 x TAE (per L): 242 g Tris base, 57.1 mL glacial acetic acid, 100 mL 0.5 M 
EDTA, pH 8.0. To prepare 1 x TAE, dilute 1:50 with distilled water. 

Acrylamide-denaturant solutions: the acrylamide solutions are only sta- 
ble for 1 month. All glassware should be rinsed with ultrapure water. 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 221 

- 8% acrylamide/0% denaturant: To make 100 mL of solution, mix 
20 mL of 40% Acrylamide/Bisacrylamide (37.5:1; BioRad); 2mL of 
50 x TAE buffer prepared with ultrapure water and ultrapure reagents, 
and 78 mL ultrapure water. Filter through a 0.22-|im filter. Mix and 
degas for 10-15 min. Store at 4°C in a brown bottle for approx. 1 
month. 

- 8% acrylamide/80% denaturant: To make 100 mL of solution mix 
20 mL of 40% acrylamide/bisacrylamide (37.5:1); 2mL of 50 x TAE 
buffer prepared with ultrapure water and ultrapure reagents; 32 mL 
deionized formamide; 33.6 g ultrapure urea, and adjust volume to 
100 mL. Filter through a 0.22- |im filter. Mix and degas for 10-15 min. 
Store at 4 °C in a brown bottle for approx. 1 month. 

• Ammonium persulfate (APS) 10% (w/v) solution: Add 100 mg of dry APS 
to 1 mL of distilled water, vortex to dissolve. This is used immediately 
and then discarded. 

• TEMED 

• Gel Loading Dye 2X (BioRad's recipe, final concentration): 0.05% bro- 
mophenol blue/0.05% xylene cyanol/70% glycerol. Prepare a 2% bro- 
mophenol blue and a 2% xylene cyanol solution. Mix 0.25 mL of each 
solution with 7.0 mL of 100% glycerol, add 2.5 mL of distilled water to 
make volume up to 10.0 mL. Store at room temperature. 

• Glycogen solution (20mg/mL; Roche Diagnostics 901393, Laval, Que., 
Canada) 

• 3 M sodium acetate (pH 5.2) 

• 100%ethanol 

• Vistra Green (Amersham Biosciences) solution: Dissolve 25 p.L of Vistra 
Green in 250 mL of lx TAE buffer (1:10,000 dilution). Store solution at 
4 °C for 3-4 days. 

• 100 bp molecular weight ladder (Fermentas SM0241) 



■ Sample Preparation 

1. PCR amplification of extracted and purified total community DNA: 
It may be necessary to dilute the soil DNA extract (preparation see 
Sect. 10.2) 1:10 or 1:100 to optimize PCR yield. 

2. A typical PCR reaction (total volume 50 pL) is composed of the following: 

- 1.0 pL of template DNA (or dilution) 



222 L.G. Whyte, C.W. Greer 

- 1.0 pL U341GC#2 primer (25 pmol); sequence: 
5^^- GCGGGCGGGGCGGGGGCACGGGGGGCGCGGCGGGC 
GGGGCGGGGG CCTACGGGAGGCAGCAG-3' (GC clamp underlined) 

- 1 .0 pL of U758 primer (25 pmol); sequence: 
5 / 758 _ 740 -CTACCAGGGTATCTAATCC-3 / 

- 0.5pLofl00mMMgCl 2 

- 8.0pLofl.25mMdNTPs 

- 32.4 pL of sterile deionized water 

- 0.625 pL of BSA (lOmg/mL; optional, but often improves the PCR 
when using DNA recovered from soils with high organic content) 

3. In a separate tube add lOx DNA polymerase buffer (5 pL per reaction) and 
DNA polymerase (0.5 pL per reaction). We typically use rTaq polymerase 
for this work. It is easier to prepare this mixture to accommodate all 
planned reaction tubes, and add 5.5 pL of the mixture to each reaction. 

4. For a "hot start" the tubes are put in the thermal cycler and heated to 
96 °C for 5 min. The temperature is then reduced to 80 °C and the DNA 
polymerase buffer/DNA polymerase mix is added to each tube. 

5. PCR is conducted using the following conditions: 

The first ten cycles use a "Touchdown protocol" from 65-55 °C, with the 
annealing temperature decreased by 1 °C at each cycle. 

- lminat94°C 

- 1 min at 65-55 °C 

- 3 min at 72 °C 

The subsequent 20 cycles are performed with an annealing temperature 
of55°C. 

7. The PCR reactions are analyzed by agarose gel electrophoresis using 
5-10 pL of reaction in a 1.4% agarose gel using TAE buffer (Sambrook 
and Russell 2001). Several dilutions (i.e., 1, 2, and 4pL of a 1:10 di- 
lution) of a 100 bp molecular weight ladder (Fermentas SM0241) are 
electrophoresed in the gel as well to quantify the amount of PCR prod- 
uct. For complex environmental samples it is advisable to prepare up to 
500 ng of PCR product to apply to each lane of the DGGE. 

■ Procedure 

Denaturant Gradient Gel (after Fortin et al. 2004) 

1. Assemble the glass plates with spacers and clamps and secure to the 
casting stand. 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 223 

2. Clamp (or tape) needle outlet from the gradient mixer between the 
glass plates (middle/top) so that it will inject the gel solution between 
the plates. 

3. To prepare a 30-70% gradient, add 7.2 mL of 8% acrylamide/0% denat- 
urant solution and 4.3 mL of 8% acrylamide/80% denaturant solution 
to one 50-mL Falcon tubes (Fisher Scientific; label "Low") and add 
1.4 mL of 8% acrylamide/0% denaturant solution and 10.1 mL of 8% 
acrylamide/80% denaturant solution to another 50 mL Falcon tubes 
(label "High"). 

4. Add 115yiL of 10% fresh APS solution to each tube. Mix gently by 
inversion. Be careful not to introduce air into the solution. 

5. Add 1 1 .5 p.L of TEMED to each tube. Mix gently by inversion. Be careful 
not to introduce air. 

6. Add the low denaturant solution gently to the left chamber (Low) of the 
gradient mixer. Remove air bubble from transfer tube by opening the 
valve stem quickly until the transfer tube between the two chambers is 
just full of low denaturant solution. 

7. Add the high denaturant solution gently to the right chamber (High) 
of the gradient mixer, turn on the mixer and the pump, open the out 
valve on the right side, and transfer the entire solution to the plates. 

8. Gently layer 1 mL of water on top of the gel to stop it from drying out. 

9. Let the gel polymerize for 1.5 h at room temperature. 

Buffer 

10. Add 6 L of 1 x TAE to gel tank (i.e., fill to the FILL line). 

11. Insert the lid and turn on. Let the buffer warm up until the temperature 
reaches 60 °C. This takes more than 1 h, so you should do this 30min 
after pouring the gel. 

Spacer Gel 

12. Using filter paper, remove the water on top of the polymerized gel. 

13. Insert gel comb fully. 

14. Mix 3.75 mL of 8% acrylamide/0% denaturant with 1.25 mL of 1 x TAE 
and with 45 iiL of 10% (w/v) APS and 4.5 \iL TEMED. 

15. Add this to the top of the denaturant gradient with a pipette. 

16. Let polymerize for 0.5 h. 



224 L.G. Whyte, C.W. Greer 

Loading and Running Gel 

17. Remove the comb and any excess polyacrylamide from the gel. 

18. Assemble the plates on the core. Pour approx. 350 mL of 1 x TAE in the 
upper chamber to check the integrity of the seal. If buffer is leaking, 
discard the buffer, disassemble, lubricate the gasket, reassemble the 
plates onto the core and test again. Insert into tank containing 1 x TAE 
buffer prewarmed to 60 °C. 

19. Let equilibrate for 15 min. 

20. Wash wells with syringe using the 1 x TAE buffer from the tank. 

21. Load wells with samples diluted in 2 x gel-loading buffer. 

22. Run gel at 80 V for 16 h at 60 °C. 



Staining Gel 

23. Stain gel for 0.5 h in 1:10,000 dilution of Vistra Green solution with 
gentle shaking. 

23. De-stain for 0.5 h in 250 mL of 1 x TAE with gentle shaking. 

24. Scan the gel on a glass plate using a Fluorlmager, save an image of the 
gel for printing (image the same size as the gel) to use as a template for 
selecting bands for excision and sequencing. 



Excising DGGE Bands and Purification for Nucleotide Sequencing 

1. Transfer the gel onto a sheet of Plexiglas under which has been placed 
the printed image of the stained gel. 

2. Cut bands of interest from gel using a scalpel or razor blade, and transfer 
into microcentrifuge tubes. 

3. Add 60 pi of sterile deionized water to each DGGE fragment and elute 
overnight in a 37 °C incubator. 

4. Centrifuge at 1 6,000 g at room temperature for 1 min and transfer super- 
natant to a fresh tube. Purify 50 pi with the QIAquick PCR purification 
kit. 

5. Add 5 volumes of Qiagen PB buffer to the 50 pi of supernatant. Vortex, 
and apply the sample to the QIAquick column-collection-tube assem- 
bly. 

6. Centrifuge at 16,000 g for 1 min (binding step). 

7. Discard flow-through. Place the column back in the collection tube. 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 225 

8. To wash, add 750 pi of Qiagen PE buffer to the column, and centrifuge 
at 16,000 g for 1 min. 

9. Discard flow- through. Place the column back in the collection tube. 

10. Centrifuge again at 16,000 g for lmin, and transfer the column to 
a 1.5 mL microcentrifuge tube. 

11. Add 50p.L of prewarmed (5 min at 50 °C) Qiagen elution buffer (EB; 
10 mM Tris-HCl, pH 8.5) to the center of the membrane. 

12. Incubate for 5 min at room temperature. 

13. Centrifuge at 16,000 g for 1 min (elution step). 

14. Store the purified DNA at -20 °C. 

15. Re-amplify the purified DNA: It is advisable to prepare 200 ng for se- 
quencing both strands. (To obtain good sequence results it is important 
to optimize PCR conditions to obtain a single band, in an agarose gel, 
of re-amplified product.) Several PCR reactions can be pooled and pre- 
cipitated: Add 1 p.1 of glycogen solution, 1/10 volume of 3M sodium 
acetate (pH 5.2) and 2.5 volumes of 100% ethanol. Precipitate at -20 °C 
for 1.5 h or overnight to accumulate sufficient product. 

16. Purify the re-amplified product using a QIAquick PCR purification kit 
or GENE CLEAN II (Qbiogene) or a GFX punification kit (Amersham 
Biosciences). 

17. Quantify the product on an agarose gel with a dilution series of a molec- 
ular standard, as described above, prior to submitting for sequencing. 

■ Notes and Points to Watch 

• The DGGE gel plates should be carefully assembled and checked for 
leaks. 

• The minimum and maximum denaturant solutions can be varied in 
concentration to change the resolution of the gel. We have found that 
a gradient from 35 to 65% denaturant gives the best resolution for many 
environmental samples. 

• When preparing denaturant solutions and adding the APS and TEMED, 
care should be taken not to introduce air. This can be accomplished by 
adding all the ingredients to separate 50-mL Falcon tubes, and gently 
mixing by inversion before adding to the gradient mixer. 

• It is important to ensure that the printed image of the scanned gel is 
identical in dimensions to the gel itself, since bands are being excised "in 



226 L.G. Whyte, C.W. Greer 

the blind" and the image is the template for removing the bands. When 
scanning the gel ensure that all four corners of the gel are scanned to 
facilitate subsequent alignment with the printed image. After removing 
the bands, the gel is re-scanned to ensure that the correct bands have 
been successfully recovered. 

10.5 

Genomics in Environmental Microbiology 

Environmental microbiology has only recently entered the genomics era. 
Genomics in its broadest sense entails the complete sequencing of an or- 
ganism's entire complement of DNA (made up of the 4 bases, A, T, C, 
and G). The sequence of DNA for a particular gene is the genetic code, 
or blueprint, that is translated into specific proteins, the key components 
in assembling all the organism's structures, regulating its functions, and 
consequently its behavior and physiology. Since the cost of genome se- 
quencing has decreased substantially, an ever increasing number of mi- 
crobial genomes are being sequenced, including important microorgan- 
isms from industrial or environmental perspectives. Many of the publicly 
available genome sequencing projects directed towards these latter or- 
ganisms are sponsored by the US Department of Energy's (DOE) Microbial 
Genome Project (www.sc.doe.gov/production/ober/microbial.html) in col- 
laboration with other partners. The DOE projects are targeting microor- 
ganisms involved in, for example, bioremediation, carbon sequestering, 
energy production, cellulose degradation biotechnology, and technology 
development. Presently 43 microorganisms with biodegradation capabili- 
ties have been/are being fully sequenced to hopefully identify new micro- 
bial processes involved in bioremediation and lead to the development of 
novel technologies and methodologies (i.e., genomic approaches, molecu- 
lar monitoring tools) for studying the structure and function of complex 
microbial communities associated with contaminated environments. 

As sequencing costs have diminished, PCR- independent methodologies 
(Fig. 10.1), including metagenomic libraries and environmental genome 
shotgun cloning approaches, have also emerged as novel ultra-high through- 
put methods to characterize complex environmental microbial communi- 
ties. Although still quite expensive, the PCR-independent methodologies 
overcome some of the limitations of the PCR-dependent methodologies, 
including the inherent bias of primer specificity in PCR amplification and 
the relatively limited amount of sequence information obtained from the 
small PCR gene targets amplified and sequenced (ca. 300- 1 ,000 nt). Metage- 
nomic libraries are created by extracting total genomic DNA from an en- 
vironment and cloning relatively large fragments (5,000-300,000 nt) into 



10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 227 

lambda, cosmid, fosmid, or bacterial artificial chromosome (BAC) vec- 
tors. The metagenomic libraries created are then screened for functional 
and/or genetic diversity that allows for clones of interest to be singled out 
and sequenced (Eyers et al. 2004). For example, Rondon et al. (2000) con- 
structed metagenomic libraries from total DNA extracted from two soils 
that contained more than 1 Gbp of DNA. Shotgun cloning is the process 
by which total community DNA is extracted from a sample, broken up to 
reduce the size, and the fragments (ca. 2,000-6,000 nt) then ligated into 
cloning vectors. Each cloning vector and the fragment of community DNA 
it carries is then amplified separately by growth in a bacterial host. The 
entire assemblage of the clone library is randomly sequenced and then 
reassembled in a procedure termed direct shotgun sequencing. Venter et 
al. (2004) performed direct shotgun sequencing (1.045 billion base pairs) 
of the microorganisms in the Sargasso Sea and identified hundreds of new 
bacterial species and 1.2 million new genes! These studies have clearly 
demonstrated the enormous biodiversity present in the environment, and 
that we have only begun to identify the vast majority of microorganisms 
out there. 

Environmental microarrays are considered an emerging technology with 
tremendous potential in the field of environmental genomics (Greer et al. 
2001; Rhee et al. 2004; Stahl 2004; Zhou and Thomson 2002). Success- 
ful application of microarray technology, which uses high-density, high- 
throughput techniques, promises to revolutionize our understanding of 
microbial diversity and microbial ecology, as thousands of potential gene 
probes can be printed on an array and hybridized to labeled total nucleic 
acids extracted from environmental samples. Environmental microarray 
technology is at a developmental stage where significant problems regard- 
ing specificity, sensitivity, and quantitation remain to be resolved (Eyers et 
al. 2004; Rhee et al. 2004; Stahl 2004). Nevertheless, application-specific 
environmental microarrays were recently used to detect sulfate-reducing 
bacteria (Loy et al. 2002), methanotrophs (Bodrossy et al. 2003), and 
biodegradative populations (Rhee et al. 2004) in environmental samples. 
Two types of environmental microarrays are presently being developed. 
Functional gene microarrays (FGMA) contain a variety of catabolic, biogeo- 
chemical cycling, heavy metal transformation genes, etc., as gene targets. 
Phylogenetic gene microarrays (PGMA) contain taxonomic gene targets, 
usually the 16S rDNA genes representing most genera of Bacteria and Ar- 
chaea. Environmental microarrays will be increasingly used to detect and 
characterize complex microbial communities in contaminated soils as well 
as to monitor degradative populations during bioremediation treatments. 
This will lead to a better understanding of important processes such as 
biogeochemical cycles and bioremediation in soils that are associated with 
mixed microbial populations in natural environments. 



228 L.G. Whyte, C.W. Greer 

Acknowledgements. The authors gratefully acknowledge the technical ex- 
pertise and contributions of Nathalie Fortin, David Juck, Diane Labbe, 
Danielle Ouellette, Sylvie Sanschagrin, Blaire Steven, Dan Speigelman, and 
Gavin Whissell. 



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11 



Bioreporter Technology 

for Monitoring Soil Bioremediation 

Steven Ripp 



11.1 

General Introduction 

Bioreporters refer to intact, living cells that have been genetically engi- 
neered to produce a measurable signal transcriptionally induced in re- 
sponse to a specific chemical or physical agent in their environment. Biore- 
porters contain three essential genetic elements, a promoter sequence, 
a regulatory gene, and a reporter gene. In the wild-type cell, the promoter 
gene is transcribed upon exposure to an inducing agent, leading to subse- 
quent transcription of downstream genes that encode for proteins that aid 
the cell in either adapting to or combating the agent to which it has been ex- 
posed. In the bioreporter, the downstream genes, or portions thereof, have 
been removed and replaced with a reporter gene. Consequently, transcrip- 
tion of the promoter gene activates the reporter gene, reporter proteins are 
produced, and some type of measurable signal is generated. These signals 
can be categorized as either colorimetric, fluorescent, luminescent, chemi- 
luminescent, electrochemical, or amperometric. Although each bioreporter 
functions differently, the end product is always the same - a measurable 
signal that is, ideally, proportional to the concentration of the specific 
chemical or physical agent to which they have been exposed (Fig. 11.1). 

Bioreporters can also be constructed without such inherent specificity. 
These bioreporters rely on reporter genes that are induced by a group of 
substances rather than just one or a few. Their primary use is for the detec- 
tion of toxic substances, which, upon exposure to the bioreporter, induce 
a stress-response gene that is fused to a reporter gene. Thus, an increase in 
signal intensity indicates toxicity, but the substance that initiated the signal 
cannot be uniquely identified. Reporter systems can also be designed to 
operate in the reverse, where a decrease in signal intensity indicates tox- 
icity. These bioreporters contain a constitutively expressed reporter gene 
that always remains on. Upon toxin exposure, the bioreporters either die or 
their metabolic activities are severely reduced, thereby causing a reduction 
in signal strength. 



Steven Ripp: The University of Tennessee, Knoxville, Tennessee, 37996, USA, E-mail: 
saripp@utk.edu 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



234 S. Ripp 



Promoter Transcription Translation 
Element q 

-^^zzza- +zz ►oj^ 




V 



I 



Reporter mRNA 

Gene Keporter 

Protein 



o ^ 

q O Inducer 

Fig. 11.1. Anatomy of a bioreporter organism. Upon exposure to a specific inducer, the 
promoter/reporter gene complex is transcribed into messenger RNA (mRNA) and then 
translated into a reporter protein that is ultimately responsible for signal generation 

Although all data generated by a bioreporter can be obtained much 
more accurately using conventional analytical techniques such as gas chro- 
matography and mass spectrometry (GC/MS), bioreporters offer a distinct 
advantage in that they report not only on a chemical's presence but on its 
bioavailability and overall effect on a living system. Bioreporters are also 
significantly cheaper, faster, and easier to use than typical analytical meth- 
ods. Additionally, for some select bioreporter systems, the bioassay can 
be performed in situ, continuously, on-line, and in real time. Such traits 
make bioreporters particularly well suited for bioremediation application. 
Typically, in any bioremediative design, the first step is to identify and 
quantify the contaminants present, which is best achieved using analytical 
techniques such as GC/MS. But after site characterization, bioreporters can 
play useful roles in the frequent monitoring that ensues, for example, when 
mapping the site to assess contaminant distribution prior to and during the 
remediation process, as well as in post-closure monitoring. Bioreporters 
can also be used to report on the status of environmental parameters impor- 
tant to successful bioremediation, such as nutrient levels, pH, and dissolved 
oxygen; or they can provide general biomass measurements (via ATP quan- 
tification) as an indication of overall microbial activity. As well, where the 
bioremediation strategy entails using an enhanced, bioengineered microor- 
ganism, the bioremediation practitioner must provide a means of tracking 
the microbe to ensure containment and monitor potential recombinant 
gene transfer events in the indigenous microbial population. This visual 
tagging can often be provided through bioreporter technology. A series of 
excellent reviews on bioreporter systems are available (Daunert et al. 2000; 
Keane et al. 2002; Belkin 2003). 



1 1 Bioreporter Technology for Monitoring Soil Bioremediation 235 

11.2 

An Overview of Reporter Systems 

for Soil Bioremediation Application 

/^-Galactosidase (lacZ) 

The lacZ gene derived from Escherichia coli encodes a /J-galactosidase 
(/J-gal) that catalyzes the hydrolysis of /J-galactosides. Traditional lacZ 
bioreporters are assayed colorimetrically. The substrate o-nitrophenyl- 
j8-D-galactoside (ONPG) is added to permeabilized bioreporter cells, af- 
ter inducer exposure, to generate a yellow by-product whose intensity 
correlates with /J-gal activity to provide an estimate of target chemi- 
cal concentration. The assay is simple and highly reliable, and has be- 
come integral to commercially available genotoxicity test kits such as 
the SOS Chromotest (Institute Pasteur, Paris). Due to low sensitivities 
and narrow dynamic ranges, however, the colorimetric test is largely 
being replaced by other detection methods. By simply using different 
/J-galactoside substrates, fluorescent, luminescent, or chemiluminescent 
assays are possible. A major disadvantage remains, however, in that the 
reporter cells must be lysed or undergo a membrane disruption step in 
order to quantify /J-gal activity. Thus, data is obtained only incremen- 
tally and results are delayed, sometimes by several hours, in relation 
to the time required to complete the /J-gal assay. Newer electrochem- 
ical and amperometric assays are beginning to solve this problem by 
measuring /J-gal activity either directly or indirectly in an on-line, near 
real-time format. However, the endogenous presence of /J-gal in natural 
environments and its potentially high background activity must always 
be taken into account when performing any of these assays. Table 11.1 
provides examples of lacZ-based bioreporters for environmental monitor- 
ing. 

Catechol 2,3-Dioxygenase (xylE) 

The xylE encoded catechol 2,3-dioxygenase is part of the pWWO plasmid 
of Pseudomonas putida and is involved in the degradation of aromatic 
compounds. Catechol 2,3-dioxygenase catalyzes the cleavage of colorless 
catechol to produce the yellow compound 2-hydroxymuconic semialde- 
hyde, forming the basis of this reporter assay. Reporter systems for xylE 
have been developed primarily for studying gene regulation, and their 
utility as environmental reporters is rather limited to the tagging of mi- 
croorganisms destined for environmental release. In this task xylE serves 
well since its endogenous activity in environmental systems is extremely 
low, as compared to lacZ. 



236 



S. Ripp 



Table 11.1. The lacZ-based bioreporters 



Analyte 


Reporter gene 


Time for induction 


Concentration 


Antimonite, arsenite 


arsR 


30min 


10 -15 M 


Biphenyls 


bphk 


3h 


ImM 


Cadmium 


zntk 


<lh 


25 nM 


Chlorocatechol 


clcR 


5 min 


10 _8 M 


Chromate 


chr 


8h 


luM 


Copper 


pcoE 


lh 


0.01 mM 




CUPl 


25 min 


0.5-2 mM 




Salmonella sulk 


2h 


0.025 pg/mL 




dink, B y D 


< 30 min 


1 pg/mL 




umuC 


3h 


0.05 pg/mL 




reck 


< 10 min 


1 pg/mL 




sfik 


2h 


< 1 ng/mL 


Mercury 


mer 


4h 


0.2 ng/mL 


Nickel 


cnr 


8h 


128 pM 


Pesticide toxicity 


HSP104 


1.5 h 


0.1 mg/L 


Phenols 


dmpR 


4h 


0.5 mM 


Zinc 


smtk 


2h 


12 pM 



^-Lactamase (bid) 

/J-Lactamase cleaves /J-lactam rings in certain antibiotics. Synthetic sub- 
strates have been developed that can also be cleaved by ^-lactamase to form 
colorimetric or fluorescent products. As with catechol 2,3-dioxygenase, 
/J-lactamase is routinely used for gene regulation studies but rarely as 
a reporter for environmental assessment. However, /Mactamase-derived 
reporters for mercury, arsenic, and cadmium are available, although their 
operational capacity under environmental conditions is unknown. 



Green Fluorescent Protein (GFP) 

Green fluorescent protein (GFP) is a photoprotein isolated and cloned 
from the jellyfish Aequorea victoria (Misteli and Spector 1997). Variants 
have also been isolated from the sea pansy Renilla reniformis. GFP pro- 
duces a blue fluorescent signal without the addition of an exogenous sub- 
strate. All that is required is an ultraviolet light source to activate the 
fluorescent properties of the photoprotein. This ability to autofluoresce 
makes GFP highly desirable in biosensing assays since it can be used 
on-line and in real time to monitor intact, living cells. Additionally, the 
ability to alter GFP to produce light emissions besides blue (i.e., cyan, 
red, and yellow) allows it to be used as a multianalyte detector. Conse- 
quently, GFP has been incorporated into bioreporters for the detection 



1 1 Bioreporter Technology for Monitoring Soil Bioremediation 



237 



Table 11.2. GFP-based bioreporters 



Analyte 


Reporter gene 


Time for induction 


Concentration 


Arsenic 


arsR 




6h 


lppb 


Benzene derivatives, 


tbu 




3h 


3.3 uM 


branched alkenes 










Biocides 


TEF 




25 min 


100 ug/mL 


Cadmium 


Cd-binding peptide 




3h 


0.5 uM 


Iron 


pvd 




Unknown 


10" 4 M 


Mercury 


mer 




16 h 


< 50 ng/mL 


Nitrate 


nar 




4h 


0.05 mM 


Octane 


alkB 




1-2.5 h 


0.01-0.1 uM 


Tetracyclines 


tetR 




50 min 


< 10 ng/mL 


Toluene 


tbuAl 




lh 


0.2 uM 


Table 11.3. Representative examples of GFP used 


as a visual tag 




Application 


Matrix 







Monitoring Arthrobacter 
Monitoring 

Pseudomonas pseudoalcaligenes 
Monitoring Alcaligenes faecalis 
Monitoring Pseudomonas fluorescens 
Monitoring Pseudomonas putida 
Monitoring Pseudomonas sp. 
Survival of Pseudomonas sp. 
Survival of Pseudomonas resinovorans 
Survival of Moraxella sp. 
Temperature effects 

on Arthrobacter chlorophenolicus 
TOL plasmid expression 
Transport of Pseudomonas putida 



4-Chlorophenol-contaminated soil 

PCB-contaminated soil 

Phenol-contaminated soil 

3-Chlorobiphenyl-contaminated root rhizosphere 

Activated sludge 

PAH -contaminated soil 

2,3-Dichlorobiphenyl-contaminatedsoil 

2,3-Dichlorodobenzo-p-dioxin-contaminatedsoil 

p-Nitrophenol-contaminated soil 

Agricultural soil 

Biofilm 

Groundwater 



of various heavy metals (Table 11.2) and as a visual tag within bacterial, 
yeast, nematode, plant, and mammalian hosts for monitoring purposes 
(Table 11.3). 



Uroporphyrinogen (Urogen) III Methyltransferase (UMT) 

UMT catalyzes a reaction that yields two fluorescent products that produce 
a red-orange fluorescence in the 590-770 nm range when illuminated with 
ultraviolet light (Sattler et al. 1995). So as with GFP, no addition of exoge- 
nous substrates is required. UMT has been used for whole-cell sensing of 
antimonite, arsenite, and arsenate (Feliciano et al. 2000). 



238 



S. Ripp 



Luciferases 

Insect Luciferase (luc). Firefly luciferase catalyzes a reaction that produces 
visible light in the 550-575 nm range. A click-beetle luciferase is also avail- 
able that produces light at a peak closer to 595 nm. Both luciferases require 
the addition of an exogenous substrate (luciferin) for the light reaction 
to occur. Examples of luc-based bioreporters constructed for the detec- 
tion of inorganic and organic compounds of environmental concern are 
presented in Table 11.4. Visual tagging of microorganisms with luc has 
also been performed in, for example, 4-chlorophenol-contaminated soils 
to track bioremediation progress. 

Bacterial Luciferase (lux). Luciferase is a generic name for an enzyme that 
catalyzes a light- emitting reaction. Luciferases can be found in bacteria, 
algae, fungi, jellyfish, insects, shrimp, and squid, and the resulting light 
that these organisms produce is termed bioluminescence. In bacteria, the 
genes responsible for the light-emitting reaction (the lux genes) have been 
isolated and used extensively in the construction of bioreporters that emit 
a blue-green light with a maximum intensity at 490 nm (Meighen 1994). 
Three variants of lux are available, one that functions at < 30 °C, another 
at < 37 °C, and a third at < 45 °C. The lux genetic system consists of five 
genes, luxA, luxB, luxC, luxD, and luxE. Depending on the combination 
of these genes used, several different types of bioluminescent bioreporters 
can be constructed. 



Table 11.4. The /wc-based bioreporters 



Analyte 


Reporter Gene 


Time for Induction 


Concentration 


Arsenite 


ars 


2h 


10 nM 


Arsenite, antimonite, 


ars 


2h 


33 nM (antimonite) 


cadmium 








Benzene, toluene, 


xylR 


30min 


3 uM (xylene) 


xylene 








Cadmium, lead, 


cadA 


2-3 h 


1 nM (antimony) 


antimony 








Chromate 


chr 


2h 


50 nM 


Copper, lead, mercury 


Drosophila Mtn 
promoter 


48 h 


3-19 ppm 


Environmental 


ERE 


10-12 h 


10 -7 M (DDT) 


estrogens 








Herbicides 


tac-luc-luxAB-aphll 


> 30 min 


ppm levels 


Mercury 


mer 


2h 


100 nM 


Organomercurials 


mer 


2h 


0.2 nM (methyl- 
mercury chloride) 


Zinc 


znt 


2h 


40 uM 



1 1 Bioreporter Technology for Monitoring Soil Bioremediation 239 

Luciferase AB (luxAB). The luxAB bioreporters contain only the luxA and 
luxB genes, which together are responsible for generating the light signal. 
However, to fully complete the light- emitting reaction, a substrate must be 
supplied to the cell. Typically, this occurs through the addition of the chem- 
ical decanal at some point during the bioassay procedure. Numerous luxAB 
bioreporters have been constructed within bacterial, yeast, insect, nema- 
tode, plant, and mammalian cell systems and have been applied toward 
detection of various environmental contaminants, monitor and control of 
bioremediation process, assays of toxicity, application of visual tags, and 
estimation of microbial biomass. 

Luciferase CDABE (luxCDABE). Instead of containing only the luxA and luxB 
genes, bioreporters can contain all five genes of the lux cassette, thereby 
allowing for a completely independent light generating system that requires 
no extraneous additions of substrate nor any excitation by an external light 
source. In these bioassays, the bioreporter is simply exposed to a target an- 
alyte and a quantitative increase in bioluminescence results, often within 
less than 1 h. Due to their rapidity and ease of use, along with the ability to 
perform the bioassay repetitively in real time and on-line, luxCDABE biore- 
porters have become extremely attractive for environmental monitoring. 
Additionally, the recent development of microluminometers for detecting 
the bioluminescent signal reduces this assay down to a miniaturized for- 
mat (Nivens et al. 2004). Table 11.5 illustrates the widespread application 
of luxCDABE-based bioreporters. 

Non-specific luxCDABE. Nonspecific lux bioreporters are typically used for 
the detection of chemical toxins. They are usually designed to continuously 
bioluminesce. Upon exposure to a chemical toxin, either the cell dies or 
its metabolic activity is retarded, leading to a decrease in bioluminescent 
light levels. Their most familiar application is in the Microtox assay (Azur 
Environmental, Newark, DE, USA) where, following a short exposure to 
several concentrations of the sample, decreased bioluminescence can be 
correlated to relative levels of toxicity (Hermens et al. 1985). The Vitotox 
test (Flemish Institute for Technological Research, Mol, Belgium) operates 
similarly (Verschaeve et al. 1999). 

Mini-Transposons as Genetic Tools in Bioreporter Constructions 

A transposon is a discrete genetic element capable of translocating from 
a donor site within the DNA molecule into one of many non-homologous 
target sites under the assistance of a transposase enzyme. Their use as re- 
porter elements was first applied in gene regulation studies using a phage 
Mu transposable element containing a promoterless lac gene. This was 
followed by similar constructs using primarily the Tn5 and TnlO family 



240 



S. Ripp 



Table 11.5. The luxCDABE-based bioreporters 



Analyte 




Reporter 


Time for induction 


Concentration 


2,3-Dichlorophenol 


reck 


(stress promoter) 


2h 


50 mg/L 


2,4,6-Trichlorophenol 


reck 


(stress promoter) 


2h 


lOmg/L 


2,4-D 




tfdRP 


20-60 min 


2 uM-5 mM 


3 -Xylene 




xyl 


hours 


3uM 


4-Chlorobenzoate 




fcbk 


lh 


380 uM-6.5 mM 


4-Nitrophenol 


reck 


(stress promoter) 


2h 


0.25 mg/L 


Anatoxin Bl 




Various stress 
promoters 


45 min 


1.2 ppm 


Ammonia 




hao 


30 min 


20 uM 


BTEX 




tod 


1-4 h 


0.03-50 mg/L 


(benzene, toluene, 










ethylbenzene, 










xylene) 










Cadmium 




cupS 


4h 


19 mg/kg 


Chlorodibromo- 


reck 


(stress promoter) 


2h 


20 mg/L 


methane 










Chloroform 


reck 


(stress promoter) 


2h 


300 mg/L 


Chromate 




chrk 


lh 


10 uM 


Cobalt 




cnr 


4-6 h 


9uM 


Copper 




Not specified 


lh 


1 uM-1 mM 


Hydrogen peroxide 




katG 


20 min 


0.1 mg/L 


Iron 




pupk 


hours 


lOnM-luM 


Isopropyl benzene 




ipb 


1-4 h 


1-100 uM 


Lead 




pbr 


4h 


4,036 mg/kg 


Mercury 




mer 


70 min 


0.025 nM 


Naphthalene 




nahG 


8-24 min 


12-120 uM 


Nickel 




cnr 


4-6 h 


0.1 uM 


Nitrate 




narG 


4h 


0.05-50 uM 


Organic peroxides 




katG 


20 min 


Not specified 


PCBs 




bph 


1-3 h 


0.8 uM 


p-Chlorobenzoic acid 




fcbk 


40 min 


0.06 g/L 


Pentachlorophenol 


reck 


(stress promoter) 


2h 


0.008 mg/L 


Phenol 


reck 


(stress promoter) 


2h 


16 mg/L 


Salicylate 




nahG 


15 min 


36 uM 


Silver 




zntkp 


lh 


0.1 uM 


Tetracycline 




tet 


50 min 


< lOng/mL 


Trichloroethylene 




tod 


1-1.5 h 


5-80 uM 


Zinc 




smtk 


4h 


0.5-4 uM 



of transposons as well as a variety of others such as Tn3/Tnl, Tn916/917, 
and TnlOOO. Although powerful mutagenic tools, natural transposons had 
several disadvantages, especially in environmental applications; they re- 
quired an antibiotic resistance marker for selection and were composed of 



1 1 Bioreporter Technology for Monitoring Soil Bioremediation 241 

inverted repeat elements that promoted unwanted genetic rearrangements 
and inherent instability (secondary transposition). They were also large 
and difficult to work with genetically and were subject to transposition im- 
munity, which prevented multiple transposon insertions within the same 
bacterial strain, severely limiting their cloning value. The development of 
mini-transposons solved many of these problems. Mini-transposons are 
shortened hybrids of natural transposons, usually Tn5 and TnlO, in which 
the transposase gene is placed outside the boundaries of the inverted re- 
peats. In this formation, the mobile element undergoes insertion into the 
target site but the transposase does not, thus preventing any further re- 
arrangements. Mini-transposons are also not affected by transposition 
immunity, thereby allowing for multiple insertions of foreign inserts in 
the same strain, provided that each insert has its own unique selectable 
marker. Additionally, mini-transposons typically maintain an origin of 
replication that allows for delivery into a broad range of hosts. Various 
mini-transposons customized with reporter genes have been developed for 
simplified construction of bioreporter organisms. By inserting a genetic 
promoter element into a unique cloning site within the mini-transposon 
vector, one can theoretically engineer any of the bioreporter classes dis- 
cussed above. Furthermore, the ability to stably insert the mini-transposon 
into the host chromosome makes these systems ideal for environmental 
applications, since the necessity for antibiotic selection can be reduced. 
Newer mini-transposons based on heavy-metal-resistance determinants 
make antibiotic selection obsolete. Methods for constructing and using 
mini-transposons are expertly described by de Lorenzo andTimmis (1994). 

11.3 

Single Point Measurements of Soil Contaminants 

■ Introduction 

Objectives. The application of bioreporters to soil bioremediation mon- 
itoring can be applied in several different formats. This can range from 
simply adding bioreporters to soil extracts to detect chemical presence to 
being so multifaceted as to use in flow-cell formats for on-line, continuous 
monitoring. With the numerous types of bioreporter systems available, the 
bioremediation practitioner has a wide range of options to choose from for 
the particular monitoring needs being addressed. For the sake of simplicity, 
the protocols described below will relate to a luxCDABE-based bioreporter 
system, but other systems can be substituted with corresponding substi- 
tutions in growth conditions, types of substrate added (if required), and 
monitoring instrumentation. The reviews by Belkin (2003), Daunert et al. 
(2000), and Keane et al. (2002) should be addressed for further direction. 



242 S. Ripp 

Principle. Bioreporter cells containing all five genes of the lux cassette are 
exposed to a soil suspension containing the target analyte, and a quantita- 
tive increase in bioluminescence is measured. 



■ Equipment 

• Centrifuge and Corex glass centrifuge tubes (Corning Inc., Corning, NY, 
USA) with Teflon screw cap lids 

• 25 mL mineralization vials with Teflon screw cap lids 

• Rotating shaker 

• Instrument capable of monitoring bioluminescence [Perkin-Elmer Vic- 
tor Multilabel reader (Wellesley, MA, USA), Azur Environmental Delta- 
tox, Zylux Femtomaster (Oak Ridge, TN, USA), Wallac Microbeta (Welles- 
ley, MA, USA), etc.] 

■ Reagents 

• YEPG medium (per L): 0.2 g yeast extract, 2 g polypeptone, 1 g glucose, 
0.2gNH 4 NO 3 ,pH7.0 

• Mineral salts medium (MSM; per L):0.1gMgSO 4 x 7H 2 O,0.2gNH 4 NO 3 , 
100 mL phosphate buffer, 0.1 mL trace elements solution 

- Phosphate buffer: 0.5 M K 2 HP04/NaH 2 P04 mixture, pH 7.0; added 
to MSM after autoclaving separately 

- Trace elements (per L distilled water): 10.0 g MgO, 2.94 g CaCl 2 , 5.4 g 
FeCl 3 x 6H 2 0, 1.44 g ZnS0 4 x 7H 2 0, 0.25 g CuS0 4 , 0.062 g H 3 B0 4 , 
0.49 g Na 2 Mo0 4 x H 2 0; added to MSM after filter sterilizing 

■ Sample Preparation 

Obtain field-moist soil samples or soil cores from test site. 

■ Procedure 

1. Preparation of bioreporter (description is for the luxCDABE bioreporter 
Pseudomonas fluorescens HK44 (Ripp et al. 2000); other bioreporter 
growth conditions will differ): 

1.1. Inoculate 100 mL of YEPG medium from a frozen stock of biore- 
porter cells. Grow overnight at 30 °C, with shaking at 200 rpm. 

1.2. The next day, inoculate 1:10 into 100 mL fresh YEPG medium. Grow 
at 30 °C with shaking to an optical density at 546 nm of 0.35. 



1 1 Bioreporter Technology for Monitoring Soil Bioremediation 243 

2. Soil preparation 

2.1. Divide soil into 10 g portions into clean 25 mLCorex glass centrifuge 
tubes. Perform in triplicate. 

2.2. Add 7 mL MSM and shake at room temperature and 200 rpm for 
lh. 

2.3. Centrifuge at 7,500 g and 25 °C for 10 min to remove large particu- 
lates. 

2.4. Remove 2 mL of supernatant into a 25 mL mineralization vial. 

3. Bioluminescent assay 

3.1. Add 2 mL of bioreporter culture (OD 546 = 0.35) to 2 mL soil extract 
in a mineralization vial. 

3.2. Transfer bioreporter/soil mixture to an appropriate holding device 
based on type of light reader being used (microtiter plate, glass vial, 
cuvette, etc.). Monitor light output for approximately 1 h. 

4. Preparation of standard curve 

4.1. Perform assay as described above with sterilized, uncontaminated 
soil to which known concentrations of contaminant have been 
added. 

5. Controls 

5.1. Perform assay as described above with sterilized, uncontaminated 
soil. Bioreporters will generate a background level of signal that 
must be subtracted from signals obtained in test samples. 

■ Calculation 

Subtract background light levels from test sample light levels and plot 
results on the standard curve to determine contaminant concentrations. 
Light levels are expressed using the arbitrary unit of relative light unit 
(RLU). 

■ Notes and Points to Watch 

• The optimal temperature for lux bioluminescent activity can be modified 
by using different lux cassettes. Vibrio fischeri lux functions at 30 °C, 
V^ harveyi at 37 °C, and Photorhabdus luminescens at 42 °C. 

• The lux reaction requires oxygen and will not operate efficiently under 
anaerobic or low oxygen conditions. 



244 S. Ripp 

• Any of the bioreporter systems described above could generate false 
positive signals due to non-specific induction by non-target inducers. 
Appropriate controls must be incorporated into experimental protocols. 

• For statistical purposes, assays should be performed in triplicate. 



11.4 

Continuous On-Line Vapor Phase Sensing 
of Soil Contaminants 

■ Introduction 

Objectives. Bioremediation processes often require a quick "snapshot" of 
soil contaminant concentrations to verify that environmental conditions are 
conducive for optimal bioremediation and to provide an overall assessment 
of where the contaminants are located. Bioreporters offer a very rapid 
assessment technology that can be economically applied to contaminated 
sites of interest. 

Principle. A flow- through chamber containing alginate-encapsulated biore- 
porter cells is inserted into a borehole and the presence of contaminants in 
the vapor phase is continuously monitored via bioluminescent signals. 

■ Equipment 

• Flow-through bioreporter chamber (Fig. 11.2): The chamber consists of 
a porous stainless steel tube (10 cm long x 2.3 cm diameter). Into the 
bottom of the tube are packed bioreporter cells encapsulated in alginate. 
Into the top of the tube is inserted a 1-mm diameter fiber-optic cable. 
The other end of the cable truncates into a photomultiplier tube [PMT; 
Hamamatsu model R-4632 Hamamatsu, Hamamatsu City, Japan), for 
example] that measures the bioluminescent signals. For complete details, 
see Ripp et al. (2000). 

• 26-gauge needle 

• Disposable 10-mL syringes 

■ Reagents 

• YEPG medium (Sect. 11.3) 

• Sterile saline solution: 0.85% NaCl 



1 1 Bioreporter Technology for Monitoring Soil Bioremediation 



245 



HK44 deriverd 
bioluminescence 



Porous 
housing 




Fiber optic 
cable 



Alginate 

encapsulated 

HK44 

bioreporter 

cells 



Fig. 1 1.2. Device for monitoring volatile PAHs using immobilized P. fluoresce ns HK44 biore- 
porters. The HK44 bioreporters bioluminesce in response to PAH exposure and the biolu- 
minescent signal is transduced through a fiber-optic cable to an external light detector 

• Sterile 3.5% (w/v) low viscosity alginic acid (Sigma- Aldrich, St. Louis) 
solution in distilled water (this must be stirred overnight, then auto- 
claved the next day) 

• Sterile 0.1 M SrCl 2 solution 

■ Sample Preparation 

Holes (approx. 4-cm diameter) need to be bored on-site, to extend into the 
zone of suspected contamination, for insertion of flow- through bioreporter 
chambers. A control hole should also be bored in a known uncontaminated 
area. 



■ Procedure 

1. Preparation of alginate-encapsulated bioreporter cells 

1.1. Grow bioreporter culture in YEPG to OD 546 = 0. 35, as described in 
Sect. 11.3. 

1.2. Centrifuge (ca. lOmin at 3,000 g) cells and wash with an equal 
volume of saline solution. Recentrifuge and again suspend in an 
equal volume of saline solution. 



246 S. Ripp 

1.3. Gently mix one part of cell suspension with two parts of alginic acid 
solution. Hold on ice. 

1.4. Gently stir 1 L of refrigerated SrCl 2 solution in a sterile 1.5-L beaker 
on a stirplate. 

1.5. Transfer the cell/ alginic acid mixture to a 10-mL disposable syringe. 
Attach a 26-gauge needle and gently push mixture through needle 
in dropwise fashion, allowing drops to fall into the beaker with the 
SrCl 2 solution. Drops will solidify into beads upon contact with 
SrCl 2 . To ensure complete solidification, allow beads to gently stir 
in SrCl 2 solution for approx. 45 min. 

1.6. Remove the beads by decanting through sterile cheesecloth. 

1.7. Store in a closed container at 4.0 °C for up to 3 months. 

2. Preparation of flow- through bioreporter chamber 

2.1. Pack 5 g of alginate-encapsulated cells into the bottom of the cham- 
ber. 

2.2. Insert fiber-optic cable directly above the encapsulated cells. 

2.3. The chamber, suspended by the fiber-optic cable, can nowbe dropped 
into the boreholes. Vapor phase detection of contaminant presence 
is then monitored continuously by the PMT and data downloaded 
to a laptop computer. Encapsulated cells must be replenished on 
a weekly basis. Newer generation sensors have reduced the bulky 
PMT modules down to an integrated circuit format, and it is now 
possible to construct wireless chip-based devices, to wit, biolumi- 
nescent bioreporter integrated circuits (BBICs), for remote sentinel 
detection of target analytes (Nivens et al. 2004). 

2.4. Background light levels should be obtained from a bioreporter 
chamber installed in a control borehole. 

■ Calculation 

Subtract light levels obtained from the control bore hole to correct for back- 
ground base levels of bioluminescence emanating from the bioreporter. 
Plot bioluminescence (RLU) versus time to illustrate trends in biolumi- 
nescence output. Based on these trends, the overall bioremediation process 
can be evaluated and monitored to determine the effectiveness of the biore- 
mediation program (i.e., low light levels will indicate unfavorable growth 
conditions for the bioremediative microbes, which should result in the 
implementation of analytical and microbiological assays and/or localized 
treatments to diagnose and correct the existing problem). 



1 1 Bioreporter Technology for Monitoring Soil Bioremediation 247 

■ Notes and Points to Watch 

• See Sect. 11.3 

• Avoid using phosphate buffers since they will degrade the alginate ma- 
trix. 

• The target contaminant(s) must produce an adequate vapor phase to be 
detected. 



11.5 

Quantification of Soil-Borne lux-lagged 
Microbial Populations Using 
Most-Probable-Number (MPN) Analysis 

■ Introduction 

Objectives. Microorganisms genetically engineered for optimal biodegra- 
dation of target contaminants can be introduced at a contaminated site to 
enhance the bioremediation process. After release, these microbes must be 
monitored to ensure that they remain within site boundaries. As well, their 
population numbers must be monitored to verify that they remain viable 
and metabolically active. The lux genes serve as excellent markers here 
because their endogenous presence in soil ecosystems is negligible, thus 
negating background interferences. The lux bioluminescent signal is also 
easily measured. The Zwx-MPN assay described below requires insertion of 
the luxCDABE cassette within the bioremediative microbe. For examples 
on how this is accomplished, see King et al. (1990). Other bioreporter sys- 
tems are applicable as well, but heightened background interferences will 
occur. For information on the introduction and release of engineered mi- 
crobes into soil and other environmental ecosystems, see Sayler and Ripp 
(2000). 

Principle. Soil inoculated with a lux-tagged bacterium is serially diluted. 
The addition of sodium salicylate results in the induction of the lux operon, 
and the intensity of bioluminescence can be correlated to cell numbers. 



Equipment 

PMT-based light reader capable of monitoring in 96-well microtiter plate 
formats [Perkin-Elmer Victor Multilabel reader, Wallac Microbeta, BMG 
Labtech Lumistar (Offenburg, Germany), etc.] 



248 S. Ripp 

■ Reagents 

• Sterile sodium pyrophosphate solution: 0.1% (w/v) 

• Sterile saline solution: 0.85% NaCl 

• Sterile sodium salicylate solution: 6 mg/mL 

■ Sample Preparation 

The contaminated soil is first inoculated with the bioremediation-enhanced, 
lux-tagged microbe. There are several methods for accomplishing this, and 
one can refer to Sayler and Ripp (2000) for general guidelines. After inocu- 
lation, soil samples (> 1 g) are removed from within areas and at depths that 
received inoculant and transported to the lab on ice. Again, P. fluorescens 
HK44 (Ripp et al. 2000) is used as an example. 

■ Procedure 

1. In a sterile test tube, add 1 g of soil to 9mL sodium pyrophosphate 
solution and vortex 1 min at top speed to remove microbes from soil 
particles. 

2. Add 100 pi of soil suspension to the first column of a 96-well black, solid 
bottom microtiter plate (Dynex Technologies, Chantilly, VA, USA). 

3. Dilute 1:2 in 100 pL saline solution throughout columns 2 through 12. 

4. Add 20 pL of sodium salicylate solution to all wells. In this example, 
sodium salicylate serves as the inducer of the lux operon in P. fluorescens 
HK44. Other bioreporters will use different inducers, but all act similarly 
to turn on the bioreporter signal. 

5. Prepare a duplicate control plate containing saline dilutions from 1 g of 
sterile soil mixed with sodium pyrophosphate solution. Add 20 pL of 
sodium salicylate solution to all wells. This plate will provide a measure 
of background bioluminescence, if any, that needs to be subtracted from 
bioluminescence counts in the sample plate. 

6. Seal plates with transparent plate sealer (Perkin-Elmer Topseal) and 
incubate at room temperature (23-28 °C) with gentle shaking for 16 h. 
This permits maximum induction of bioluminescence from HK44 cells. 
Use of other bioreporters will require optimization of incubation times. 

7. Measure photon emission from wells with a microtiter-based light reader, 
such as the Perkin-Elmer Victor instrument. Read each plate in triplicate 
for statistical verification. 



1 1 Bioreporter Technology for Monitoring Soil Bioremediation 249 

■ Calculation 

Input data into any variety of MPN software programs (see, for example, 
Klee 1993). These programs will use Poisson statistics to estimate cell 
numbers based on where bioluminescence is first observed within the 
dilution series. 

■ Notes and Points to Watch 

• See Sect. 11.3. 

• Accurate MPN population estimates require accurate dilutions. Ensure 
that all dilution series are performed as carefully as possible. 

References 

Belkin S (2003) Microbial whole-cell sensing systems of environmental pollutants. Curr 
Opin Microbiol 6:206-212 

Daunert S, Barrett G, Feliciano JS, Shetty RS, Shrestha S, Smith-Spencer W (2000) Genetically 
engineered whole-cell sensing systems: coupling biological recognition with reporter 
genes. Chem Rev 100:2705-2738 

de Lorenzo V, Timmis KN (1994) Analysis and construction of stable phenotypes in gram- 
negative bacteria with Tn5- and TnlO-derived minitransposons. Methods Enzymol 
235:386-405 

Feliciano J, Liu Y, Ramanathan S, Daunert S (2000) Fluorescence-based sensing system for 
antimonite and arsenite using cob A as the reporter gene. 219th ACS National Meeting 

Hermens J, Busser F, Leeuwangh P, Musch A (1985) Quantitative structure-activity relation- 
ships and mixture toxicity of organic chemicals in Photobacterium phosphoreum: the 
Microtox test. Ecotoxicol Environ Saf 9:17-25 

Keane A, Phoenix P, Ghoshal S, Lau PCK (2002) Exposing culprit organic pollutants: a review. 
J Microbiol Meth 49:103-119 

King JMH, DiGrazia PM, Applegate B, Burlage R, Sanseverino J, Dunbar P, Larimer F, 
Sayler GS (1990) Rapid, sensitive bioluminescence reporter technology for naphthalene 
exposure and biodegradation. Science 249:778-781 

Klee AJ (1993) A computer program for the determination of most probable number and 
its confidence limits. J Microbiol Meth 18:91-98 

Meighen EA (1994) Genetics of bacterial bioluminescence. Annu Rev Genet 28:117-139 

Misteli T, Spector DL (1997) Application of the green fluorescent protein in cell biology and 
biotechnology. Nat Biotechnol 15:961-964 

Nivens DE, McKnight TE, Moser SA, Osbourn SJ, Simpson ML, Sayler GS (2004) Biolu- 
minescent bioreporter integrated circuits: potentially small, rugged and inexpensive 
whole-cell biosensors for remote environmental monitoring. J Appl Microbiol 96:33-46 

Ripp S, Nivens DE, Ahn Y, Werner C, Jarrell J, Easter JP, Cox CD, Burlage RS, Sayler GS (2000) 
Controlled field release of a bioluminescent genetically engineered microorganism for 
bioremediation process monitoring and control. Environ Sci Technol 34:846-853 

Sattler I, Roessner CA, Stolowich NJ, Hardin SH, Harris-Haller LW, Yokubaitis NT, 
Murooka Y, Hashimoto Y, Scott AI (1995) Cloning, sequencing, and expression of the 
uroporphyrinogen-III methyltransferase cob A gene ofPropionibacteriumfreudenreichii 
(shermanii). J Bacterid 177:1564-1569 



250 S. Ripp 

Sayler GS, Ripp S (2000) Field applications of genetically engineered microorganisms for 

bioremediation processes. Curr Opin Biotechnol 11:286-289 
Verschaeve L, Van Gompel J, Thilemans L, Regniers L, Vanparys P, van der Lelie D (1999) 

VITOTOX bacterial genotoxicity and toxicity test for the rapid screening of chemicals. 

Environ Mol Mutagen 33:240-248 



^ ^ Interpretation of Fatty Acid Profiles 
' ^ of Soil Microorganisms 

David B. Hedrick, Aaron Peacock, David C. White 



12.1 

Obtaining Fatty Acid Profiles from Soil Samples 

This work focuses on the calculations performed on the peak areas obtained 
by gas chromatography (GC). All the steps of soil sampling, lipid extraction 
and fractionation, derivatization, and capillary GC have been repeatedly re- 
viewed, and will only be briefly mentioned (a bibliography of work done in 
this laboratory is available at http://cba.bio.utk.edu/director_peerfull.html, 
and an extensive bibliography of methods is provided by Dr. William 
Christie's group, Mylnefield Research Services Ltd. at http://www. 
lipidlibrary.co.uk/lit_surv.html). 

Sampling is the most important step in sample analysis, and is often 
delegated to the most junior member of the lab or to site specialists not 
associated with the lipid laboratory, such as a subsurface sediment drilling 
crew. Besides sampling location, the sample's consistency, integrity, and 
appearance should be recorded. In order to obtain deep subsurface sam- 
ples, the use of drilling equipment and drilling mud is usually required, 
and methods have been developed to prevent and detect drilling mud con- 
tamination of samples (Griffin et al. 1997; Phelps et al. 1989). 

Capillary GC with flame ionization detection (FID) is a powerful ana- 
lytical method - simpler in operation, of greater linear range, and more 
sensitive, reliable, and reproducible than most analytical instrumentation 
available. The users' manuals for the chromatograph and data system are 
the primary references for their operation. If you won't read the manual, 
you shouldn't touch the equipment. There are also many excellent reviews 
of capillary chromatography of polar lipid fatty acids (PLFA) available (for 
example, Grob and Barry 1995). 

Capillary GC-MS is a necessary adjunct to GC-FID for the identification 
of fatty acid peaks (Christie 2003). Various chemical methods are also avail- 
able to help with specific identification problems such as silver ion chro- 
matography to separate saturates, monounsaturates, and polyunsaturates 



David B. Hedrick, Aaron Peacock, David C. White: Center for Biomarker Analysis, University 
of Tennessee, 10515 Research Drive, Suite 300, Knoxville, Tennessee 37932-2575, USA, 
E-mail: dwhitel@utk.edu 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



252 D.B. Hedrick et al. 

(Momchilova and Nikolova-Damyanova 2000), and special derivatization 
methods to determine the position and geometry of monounsaturation, 
such as MS of dimethyldisulfide adducts (Nichols et al. 1986). MS of picol- 
inyl esters provides more informative fragmentations than GC-MS of the 
methyl ester (Christie et al. 1991; Harvey 1992). 

This work presupposes some knowledge of Microsoft Excel (Microsoft 
Corp., Redmond, WA), which is used to manipulate chromatographic re- 
sults in many laboratories. The on-line help system is the basic reference 
for Excel, such as it is. A novice user will benefit from one of the many intro- 
ductory books available at a bookstore. Also assumed is some background 
in the statistical procedures commonly applied to PLFA data, including 
analysis of variance (ANOVA) and factor analysis. 

12.2 

Transforming Fatty Acid Peak Areas 

to Total Microbial Biomass 

Gas chromatography provides a peak area proportional to the amount of the 
compound in the sample responsible for the peak. A known concentration 
of an internal standard, usually 19:0 or 21:0, is added to the sample before 
analysis to allow calculation of absolute amounts (see Sect. 12.5 for the 
naming of fatty acids). The equation used to calculate the total amount of 
fatty acids in a sample is, 

(sum A F a/Ats) x IS x X , 

FA=- ^— — 12.1 

Y 

FA total picomoles of fatty acids per gram dry mass of sample (pmol/g 

dry mass) 

sum A F a sum of the areas of all identified fatty acid peaks excluding the 
internal standard 

A IS area of the internal standard peak 

IS concentration of internal standard used (50 pmole/p.L) 

X volume of internal standard used to dilute the fatty acid methyl 

esters (p.L) 

Y mass of sample extracted (g soil dry mass). In some instances, 

rather than grams dry mass as the divisor, it will be volume of 
water (L), surface area in meters squared, or some other extensive 
variable. 



12 Interpretation of Fatty Acid Profiles of Soil Microorganisms 253 

Many analysts calculate the pmol/g dry mass for each fatty acid, then 
add them together to get the total pmole/g dry mass. This is not good 
practice, since the pmol/g dry mass for each fatty acid is not then of use in 
further analysis, and the more complicated calculation makes more work 
and opportunities for error. 

The total moles of membrane fatty acids is proportional to the total 
microbial biomass. The constant of proportionality used in our laboratory 
is 2.5 x 10 4 cells/pmol PLFA (Balkwill et al. 1988; White et al. 1996 and 
references therein). This conversion factor was derived from measurements 
on laboratory cultures, so the number of cells will be underestimated for 
environments populated by smaller bacterial cells, such as oligotrophic 
environments. 

Researchers who count cells, with automated cell counting instruments 
or by microscopy, are often uncomfortable with measurements of viable 
biomass expressed as moles of PLFA or grams dry mass of cells. In order 
to estimate cell counts from moles of PLFA requires knowledge of the 
distribution of cell sizes in the sample and the amount of PLFA per cell for 
different sizes, information which is not usually available. It makes more 
sense to transform cell counts to moles PLFA or from the latter to grams 
dry weight of cells, since the cell counting can provide the data on cell size 
distribution. 

For most sample sets, the biomass will not be normally distributed, that 
is, a histogram of the biomass data will be skewed with a long tail toward 
the higher biomasses. This can be tested for by using the standard f-test 
for normality. Also, in most biomass data sets, the variance of biomass 
increases with the absolute value of the biomass. This violates the assump- 
tions of parametric statistics, including ANOVA and factor analysis, and 
lowers the power of any statistical test employed. These problems can be 
solved by a log(X + A) transformation, where X is the mole percent of the 
fatty acid, and A is a small constant. The small constant is added so that 
zero values give a real solution when the log transform is applied. The 
most common value used for A is one, which gives a value of zero for the 
transform when X is zero, since log(0 + 1) = 0. 

There are two approaches to proving the value of applying a log trans- 
form to biomass data, the theoretical and the practical. The theoretical 
explanation involves the scaling of the forces affecting microbial biomass 
(Magurran 1988) and the fractal structure of microbial environments (Man- 
delbrot 1982), and is beyond the scope of this work. The practical reason 
for the log transform is that it works; applying a log transformation to the 
data is perfectly legitimate, and results in more significant differences on 
statistical tests. 



254 



D.B. Hedrick et al. 



12.3 

Calculation and Interpretation of Community Structure 

After the biomass, the next most important information to extract from 
a PLFA profile is the community structure. But where the biomass is a single 
value for each sample with a straightforward interpretation, the commu- 
nity structure data is multivariate with many options in its interpretation. 
A "standard" method for presenting community structure data, how to 
create a custom method for community structure, and factor analysis will 
be presented. 



12.3.1 

Standard Community Structure Method 

In the standard method for community structure analysis of PLFA pro- 
files, chemically related fatty acids are grouped as in Table 12.1. A PLFA 
profile may contain, for example, from 18 to 92 fatty acids. The standard 
community structure approach summarizes that in six variables, which are 
just the sum of the mole percents of each of the fatty acid groups. The use 
of a standard community structure analysis method allows comparison 
between/among experiments. 



Table 1 2. 1 . Groups of chemically related fatty acids used in the standard community structure 
analysis 



Group name 


Rule 


Examples 


Microbiota represented 


Saturates 


Saturated straight- 


12:0, 13:0, 14:0, 


All organisms 




chain fatty acids 


15:0, 16:0, 17:0, 
18:0 




Monounsaturates 


Fatty acids with 


14:lcc>5c, 


Proteobacteria 




a single unsaturation 


16:1g)7c, 






plus cyclopropyls 


I6:loo7t, 
\S:lco7c 




Mid-chain branched 


Any mid- chain 


10Mel6:0, 


Actinomycetes, 




branched fatty acid 


10Mel8:0 


sulfate-reducers 


Terminally branched 


Iso- and anti-iso- 


il4:0,il5:0, 


Gram positive bacteria 




branched saturated 


al5:0,il6:0, 






fatty acids 


il7:0,al7:0 




Polyunsaturates 


Any fatty acid with 


18:2a)6c, 


Eukaryotes 




more than one 


1S:3co3c 






unsaturation 






Branched unsaturates 


Any branched 
monounsaturate 


H7:lco7c 


Anaerobes 



12 Interpretation of Fatty Acid Profiles of Soil Microorganisms 255 

The standard community structure breakdown was originally devel- 
oped on marine sediments, and has been successfully applied to microbial 
communities from many environments, including, for example, marine 
macrofaunal burrows (Marinelli et al. 2002), a subsurface zero-valent iron 
reactive barrier for bioremediation (Gu et al. 2002), marine gas hydrates 
(Zhang et al. 2002), soils contaminated with jet fuel (Stephen et al. 1999), 
and to a comparison of subsurface environments (Kieft et al. 1997). 

12.3.2 

Custom Community Structure Methods 

When examination of the chromatograms or the mole percent table shows 
differences with treatment, but no significant differences are found in the 
standard community structure groups, some other way of grouping the fatty 
acids maybe more useful. For example, if samples differ in the proportions 
of Cyanobacteria and Eukaryotic algae, it may be useful to separate the 
polyunsaturates with 18 or fewer carbons characteristic of Cyanobacteria 
(0ezanka et al. 2003) from those typical of Eukaryotic algae with 20 or 
more carbons (Erwin 1973). 

There are several methods for developing alternative community struc- 
ture groups. The manual method uses the pattern recognition power of the 
human eye. The PLFA chromatograms are printed on the same scale and 
spread out on a large table. Similar-looking chromatograms are grouped 
together and different-looking ones are placed in separate groups. While 
very low- tech, this works remarkably well. This same approach can be ap- 
plied to a mole percent table by printing it out, cutting out a strip for each 
sample, and sorting the samples by similarity. Once the samples have been 
sorted into similar groups, the fatty acids responsible are summed to form 
new community structure groups. 

Given access to statistical software, a triangular table of Pearson's r 
correlation coefficients is usually available as an output option. Visual 
examination of this table will locate fatty acids with high correlations, which 
are then grouped together to form new community structure groups. 

12.3.3 

Factor Analysis 

Factor analysis includes several related methods, including principal-com- 
ponents analysis. The virtue of this method is that it automatically con- 
structs fatty acid groups reflecting the differences in community structure, 
rather than applying a preconception of fatty acid groups. The data deter- 
mines the fatty acid groups, rather than the analyst. Factor loadings greater 



256 D.B. Hedrick et al. 

than 0.7 indicate fatty acids with "significant" effects on the results. The 
factor scores are new variables that are linear combinations of the origi- 
nal values. These new variables can be submitted to statistical tests such 
as ANOVA like any other variable. Examples of the application of factor 
analysis to PLFA profiles include storage perturbation of soil microbial 
communities (Haldeman et al. 1995; Brockman et al. 1997), soils at differ- 
ent temperatures (Zogg et al. 1997), and soils from different ecosystems 
(Myers etal. 2001). 

The results of factor analysis are usually improved by applying the log(X+ 
1) transformation to the mole percent data before factor analysis. A rough 
method to determine whether the mole percent data is normally distributed 
is to calculate the maximum, average, and the minimum not equal to zero 
for each fatty acid. The formulas for these in Excel are "= max(b2.b45)", "= 
average(b2.b45)", and "= min(if(b2.b45 = 0, 100,b2.b45))", where b2.b45 
is the range containing the data. The formula for min is what Excel 
terms an array formula; you have to hold down the Shift and Control keys 
while you press Enter to enter the formula. If the difference between the 
maximum and average is greater than the difference between the average 
and the minimum for most of the fatty acids, then the data is not normally 
distributed and the log(X + 1 ) transformation will probably improve results. 

There are theoretical reasons to advocate the arcsin[square root(X)] 
transformation over the log(X + 1 ) transformation, but very little difference 
is found in practice, and the log(X + 1) is simpler to apply and explain. 
Similarly, there are theoretical reasons to prefer factor analysis sensu stricto 
over principal components analysis, and vice versa, which can, and have 
been, argued for days to no conclusion. In practice, the two methods give 
very similar results. 

12.4 

Calculation and Interpretation 

of Metabolic Stress Biomarkers 

The membrane of the bacterial cell handles all of its interactions with 
its environment, and bacteria have many strategies to deal with stressful 
environmental conditions, including modifying the fatty acids used in the 
membrane. This is illustrated in Eq. (12.2), where S stands for the substrate 
fatty acid and P for the product fatty acid induced by metabolic stress, 
namely, a trans monounsaturate or cyclopropyl fatty acid. 

S^ P 

cis monounsaturate — >► trans monounsaturate (12.2) 

cis monounsaturate —> cyclopropyl 



12 Interpretation of Fatty Acid Profiles of Soil Microorganisms 257 

The stress biomarkers are then calculated as the ratio of the mole percents 
of the product to the substrate fatty acids, as in Eq. (12.3): 

BMs tr ess = P/S (12.3) 

where BM Str ess is the value of the stress biomarker. The most common trans- 
formations are !6:lco7c^ \6:lco7 t, 16:l(x>7c^Cyl7:0, lS:lco7c^ 18:loo7 t, 
andl8:lco7c^Cyl9:0. 

There are problems with the application of the stress biomarkers. The 
first type of problem is when the stress-induced product fatty acid is only 
detected in a minority of the samples. This will most likely prevent detection 
of statistically significant differences. The second problem is when the 
substrate fatty acid is not detected, but the stress-induced fatty acid is; this 
has been seen in hot acid environments such as hydrothermal systems. 
Since division by zero is undefined in standard algebra, undefined results 
appear that standard statistical programs are unable to use. This problem 
can be solved by a modification of Eq. (12.3), 

BMs tr ess = P/(S + 1) (12.4) 

The metabolic stress biomarkers have been applied to, for example, tap 
water biofilms (White et al. 1999), and soils contaminated with jet fuel 
(Stephen et al. 1999). 



12.5 

Naming of Fatty Acids 

Creating clear, consistent, and unambiguous names for microbial fatty acids 
is challenging due to the wide variety of possible structures. At the same 
time, it is essential for understanding the data and communicating results. 
The IUPAC rules for naming chemical compounds are supposed to provide 
unambiguous names, but there are problems with this approach. The most 
important is that IUPAC counts carbons from the opposite end of the fatty 
acid molecule from most of the enzymes that modify the fatty acid. 

The need for a compact notation has led to the development of the 
omega system for naming fatty acids. Fatty acids are named according to 
the pattern of AiBcoC. The A stands for the number of carbon atoms in the 
fatty acid backbone, B is the number of double bonds, and C is distance 
of the nearest unsaturation from the aliphatic (co) end of the molecule. 
This can be followed by a "c" for cis or a "t" for trans configuration of 
the unsaturation. The prefixes "i," "a," and "br" stand for iso, anti-iso, 
and unknown branching position of the carbon chain, respectively. Mid- 
chain branching is noted by a prefix "lOMe" for a 10-methyl fatty acid, and 



258 D.B. Hedrick et al. 

cyclopropyl fatty acids by prefix "Cy." For example: \S:lco7c is 18 carbons 
long with one double bond occurring at the 7th carbon atom from the co 
end, and the unsaturation is in the cis conformation. Also, 16:0, il6:0, al6:0, 
and brl6:0 are all 16-carbon fatty acids, while 10Mel6:0 and Cyl7:0 both 
contain a total of 17 carbons, not counting the carbon of the methyl ester 
moiety. 



References 

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bial biomass measures based on membrane lipid and cell wall components, adenosine 
triphosphate, and direct counts in subsurface sediments. Microbial Ecol 16:73-84 

Brockman FJ, Li SW, Fredrickson JK, Ringelberg DB, Kieft TL, Spadoni CS, White DC, 
McKinley JP (1997) Post-sampling changes in microbial community composition and 
activity in a subsurface paleosol. Microbial Ecol 36:152-164 

Christie WW (2003) Lipid analysis; isolation, separation, identification and structural anal- 
ysis of lipids, 3rd edn. Oily Press, Bridgwater, UK 

Christie WW, Brechany EY, Lie Ken Jie MSF, Bakare O (1991) MS characterization of picolinyl 
and methyl ester derivatives of isomeric thia fatty acids. Biol Mass Spectrom 20:629-635 

Erwin JA (1973) Fatty acids in eukaryotic microorganisms. In: Erwin JA (ed) Lipids and 
biomembranes of eukaryotic microorganisms. New York, Academic Press, pp 41-143 

Griffin WT, Phelps TJ, Colwell FS, Fredrickson JK (1997) Methods for obtaining deep 
subsurface microbiological samples by drilling. In: Amy PS and Haldeman DL (eds) 
The microbiology of the terrestrial and deep subsurface. CRC Press, Boca Raton, pp 23- 
43 

Grob RL Barry EF (1995) Modern practice of gas chromatography. Wiley, New York 

Gu B, Zhou J-Z, Watson DB, Philips DH, Wu L, White DC (2001) Microbiological character- 
ization in a zero-valent iron reactive barrier. Appl Environ Microbiol 77:293-309 

Haldeman DL, Amy PS, Ringelberg DB, White DC (1994) Changes in bacteria recoverable 
from subsurface volcanic rock samples during storage at 4 °C. Appl Environ Microbiol 
60:2679-2703 

Harvey DJ (1992) Mass spectrometry of picolinyl and other nitrogen-containing derivatives 
of fatty acids. In: Christie WW (ed) Advances in lipid methodology, vol 1. Oily Press, 
Ayr, UK, pp 19-80 

Kieft TL, Murphy EM, Amy PS, Haldeman DL, Ringelberg DB, White DC (1997) Laboratory 
and field evidence for long-term starvation survival of microorganisms in subsurface 
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vestigation of extraterrestrial organisms, 27 July to 1 August. Int Soc Optical Engin, San 
Diego, CA 

Magurran AE (1988) Chapt. 2. Diversity indices and species abundance models. In: Magur- 
ran AE (ed) Ecological diversity and its measurement. Princeton Univ Press, Princeton, 
NJ 

Mandelbrot B (1982) The fractal geometry of nature. Freeman, San Francisco, CA 

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vestigation of the control of bacterial community composition in macrofaunal burrows. 
Marine Ecol Prog Series 235:1-13 

Momchilova S, Nikolova-Damyanova B (2003) Stationary phases for silver ion chromatog- 
raphy of lipids: Preparation and properties. J Sep Sci 26:261-270 



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community composition and substrate, use in forest ecosystems. Soil Sci Soc Am J 
65:359-367 

Nichols PD, Guckert JB, White DC (1986) Determination of monounsaturated double bond 
position and geometry for microbial monocultures and complex consortia by capillary 
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0ezanka T, Dor I, Prell A, Dembitsky VM (2003) Fatty acid composition of six freshwater 
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Stephen JR, Chang Y-J, Gan YD, Peacock A, Pfiffner SM, Barcelona MJ, White DC, Mac- 
naughton SJ (1999) Microbial characterization of JP-4 fuel contaminated-site using 
a combined lipid biomarker/PCR-DGGE based approach. Environ Microbiol 1:231-241 

White DC, Kirkegaard RD, Palmer Jr. RJ, Flemming CA, Chen G, Leung KT, Phiefer CB, 
Arrage AA (1999) The biofilm ecology of microbial biofouling, biocide resistance and 
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Soc Amer J 61:475-481 



13 



Enumeration of Soil Microorganisms 

Julia Foght, Jackie Aislabie 



13.1 

Sample Preparation and Dilution 

■ Introduction 

Objectives. Soil is a heterogeneous matrix in which microbes are associated 
with organic and inorganic soil particles, forming aggregates. The goals of 
sample preparation for conventional enumeration techniques are to release 
the microbes from the matrix of a representative soil sample, then disperse 
them in a suitable diluent so that individual cells can be enumerated ei- 
ther by microscopic visualization or cultivation methods. The basic meth- 
ods for soil aggregate disruption and dilution have been in common use 
for decades, but individual laboratories often develop variations to create 
their own empirical "standard methods." Different soil types may be more 
amenable to certain diluents or disruption techniques, so, if examining an 
unfamiliar soil type, it is wise to test combinations of methods to empir- 
ically optimize enumeration results. The presence of inorganic or organic 
contaminants (e.g., crude oil) may require adaptation of the basic methods 
to disperse the soil sample adequately or dilute a toxicant (e.g., heavy metal). 

Principle. A suitable buffered diluent releases microbial cells from the soil 
matrix and is used to dilute the suspension to a cell density suitable for the 
enumeration method to be used. The dilution method must not compro- 
mise the structural integrity of cells to be enumerated by microscopy, nor 
the viability of cells for culture-based enumeration. 

Theory. Microbes in soil are distributed heterogeneously in microenviron- 
ments of different scales and along depth profiles (Foster 1988; Ranjard and 
Richaume 2001). Therefore, representative samples of a suitable size must 
be collected for accurate enumeration. The number of individual samples 
theoretically required to represent the site can be calculated (Alef and Nan- 
nip ieri 1995), but in practical terms the number of samples handled is 



Julia Foght: Biological Sciences, University of Alberta, Edmonton AB, Canada T6G 2E9, 
E-mail: julia.foght@ualberta.ca 

Jackie Aislabie: Landcare Research, Private Bag 3127, Hamilton, New Zealand 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



262 J. Foght, J. Aislabie 

dictated by the time and resources available. As a compromise, a composite 
sample can be prepared from several samples of equal mass or volume, but 
statistical evaluation of the data is relinquished. Commonly, at least 10 g wet 
mass of soil is used to prepare the first dilution, although the sample size 
maybe adjusted according to the soil type and the organisms to be enumer- 
ated. Serial dilutions (commonly ten-fold) of soil suspensions are prepared 
with sufficient mixing to disrupt soil aggregates and release occluded mi- 
crobes into suspension. Physical disruption of the soil aggregates can be 
enhanced by inclusion of small (2-3 mm) sterile glass beads in the diluent, 
at least in the first dilution. Suitable sterile diluents, of which many exist, 
aid the dispersion of soil aggregates. Diluents are often buffered (Strick- 
land et al. 1988) and may contain proteins such as gelatin or tryptone to aid 
dispersion, glycerol to aid resuscitation of starved bacterial cells (Trevors 
and Cook 1992), or a surface active agent such as 0.1% Tween 80, although 
surfactants may reduce counts of sensitive Gram-negative cells (Koch 1994). 

■ Equipment 

• Top-loading balance capable of weighing to 0.1 g 

• 150-mL glass dilution bottles and, optionally, approx. 20 g of 2-3 mm 
glass beads per bottle to aid in disruption of soil aggregates 

• Spatula or small spoon, sterilized by autoclave or by flaming with ethanol 

• Sterile pipettes for serial dilutions: 10-mL wide-mouth glass pipettes are 
less likely to plug during initial dilutions 

• Optional mixing equipment: reciprocating or gyratory shaker for first 
dilution; vortex mixer; Waring blender 

■ Reagents 

• Suitable sterile, buffered diluent dispensed into dilution bottles, usually 
90 or 99 mL each 

• Suitable diluents include: 0.1% (w/v) sodium pyrophosphate with or 
without 1% glycerol (Trevors and Cook 1992); phosphate-buffered saline 
(0.85% (w/v) NaCl, 2.2 mM KH 2 P0 4 ; 4.2 mM Na 2 HP0 4 , pH 7) with or 
without 0.01% gelatin or peptone (Koch 1994); 1-10 mM potassium 
phosphate (pH 7); or mineral salts medium lacking carbon source (Atlas 
1995). 

■ Sample Collection 

Acceptable aseptic techniques for collection and storage of soil samples are 
given in Chapt. 1 in this volume. Soil intended for conventional enumeration 



1 3 Enumeration of Soil Microorganisms 263 

techniques should not be dried because this can reduce the microbial counts 
(Sparling and Cheshire 1979; van Elsas et al. 2002). Analyses should be 
conducted as soon as possible after sample collection. 

■ Procedure 

1. On a top-loading balance use sterile spatula to aseptically dispense 10 g 
of soil into the first dilution bottle containing 90 mL of diluent and 
record exact wet mass of sample added. This is the 10 _1 dilution. Alef 
and Nannipieri (1995) recommend using 20 g soil in 180mL of diluent 
to reduce the effects of sample heterogeneity. 

2. To express the counts on the basis of soil dry mass, dispense a similar 
sample into a tared aluminum pan for determining dry mass (in triplicate 
for accuracy). Dry the sample at 105 °C to constant mass overnight, and 
record mass. 

3. Shake or mix the dilution bottle vigorously manually or mechanically 
(using reciprocating shaker or Waring blender) to disrupt soil aggre- 
gates; recommended times vary from 1 min to 1 h and can be optimized 
empirically for different soils. 

4. Perform ten-fold dilutions by transferring a 10.0-mL sample from the 
center of the dilution bottle to a fresh 90-mL dilution bottle, or hundred- 
fold dilutions with 1.0 mL transferred into 99 mL of diluent. Mixing 
between dilutions may be performed by hand by vigorously shaking the 
bottle 25 times between each transfer, or with a vortex mixer. 

5. Continue with ten-fold serial dilutions appropriate to the enumeration 
method to be used, e.g., for aerobic heterotrophs in uncontaminated 
agricultural soils dilute to 10" 9 for most probable number (Sect. 13.3) 
and 10~ 7 for plate counts (Sect. 13.4). 

■ Calculation 

1. Dilution factor (reciprocal of dilution) = (1/dilution) 

2. Dry-mass correction factor = (wet mass of sample/dry mass of sample) 

■ Notes and Points to Watch 

• The initial sample(s) must be as representative of the soil as possible and 
analysis of replicates is recommended. 

• Sample preparation and dilutions must be performed in a standardized 
manner that can be replicated, so that results from samples taken at 



264 J. Foght, J. Aislabie 

different times or from different sample sites can be compared with 
confidence. 

• Soil dilutions should be used immediately after preparation, as storage 
of the cell suspension in buffer may decrease the counts observed (Koch 
1994). 

• The dilution volumes can be scaled down, using test tubes with 1 g of 
soil in 9 mL of diluent and mixing by vortex, but caution should be used 
because small sample sizes may not be representative. 

• A sonicator bath or probe may be used for initial soil sample disrup- 
tion (Strickland et al. 1988), but this equipment is not standard in all 
laboratories, and excess sonication will reduce counts. 

• Aggregates in hydrocarbon-contaminated soils may be difficult to dis- 
perse, yielding inaccurate results. Similarly, microbes with highly hy- 
drophobic cell surfaces, such as acid-fast hydrocarbon-degrading bacte- 
ria, may themselves aggregate and be difficult to disperse. 

• If using sodium pyrophosphate as the diluent, adjust the pH to neutrality, 
as it is ca. pH 10 without adjustment (Trevors and Cook 1992). 



13.2 

Direct (Microscopic) Enumeration 

■ Introduction 

Objectives. It has long been known that enumeration techniques relying on 
cultivation of microbes in environmental samples can underestimate the 
total number of cells present by orders of magnitude (Skinner et al. 1952; 
Amann et al. 1995). This bias can be overcome in part by using molecular 
methods (Chapt. 10) or by using direct microscopic observation of cells 
where no cultivation is required. Direct enumeration methods can provide 
the total number of cells (live plus dead) or may discriminate between live 
and dead cells. Some stains differentiate cells based on phylogeny or the 
presence of functional genes, providing information about the types of cells 
as well as numbers. Microscopy is suitable for direct enumeration of both 
bacteria and fungi. 

Principle. A known volume of a soil suspension is filtered through a 0.2 p.m 
pore size filter. The microbes on the filter are stained with a fluorescent dye 
and counted by using an epifluorescence microscope. At least 20 fields each 
containing 20-50 cells are counted and the total count is calculated from 
the area observed and the volume of suspension filtered. 



1 3 Enumeration of Soil Microorganisms 265 

Theory. To reduce the bias inherent in culture-based enumeration meth- 
ods, total counts of microbes in soil can be observed directly using mi- 
croscopy (Fry 1990; Kepner and Pratt 1994; Bottomley 1994; Bloem 1995). 
Traditionally, to aid detection, the cells have been stained with fluores- 
cent dyes (reviewed by Bolter et al. 2002) such as acridine orange (AO) 
or 4 / ,6-diamino-2-phenylindole (DAPI) which stain DNA- containing cells. 
Recently, emphasis has been put on differentiating between actively metab- 
olizing cells and resting cells, or on discriminating between live and dead 
cells. Hence, new fluorescent dyes have been developed. The redox dye 5- 
cyano-2,3-ditolyl tetrazolium chloride (CTC), for example, is used to count 
active bacterial cells (Creach et al. 2003). CTC is a colorless membrane- 
permeable compound that produces a red-fluorescing precipitate in the 
cell wall when reduced by the electron transport system of active bac- 
terial cells. Staining with a combination of propidium iodide (PI, which 
is excluded from cells with intact membranes) and thiazole orange (TO, 
which is taken up by both live and dead cells) provides a method for dis- 
criminating between live and dead cells. Numerous commercial stain kits 
are available with specific instructions for their use, such as Live/Dead 
BacLight kits (Molecular Probes, Invitrogen, Carlsbad, CA, USA). The flu- 
orescent in situ hybridization (FISH) method, which detects hybridization 
of fluorescently-labeled oligonucleotide probes with target DNA or RNA 
sequences, can combine total counts with counts of specific phylogenetic 
groups (Amman et al. 1995) by detecting multiple overlapping fluorescent 
signals, but, like other microscopic methods, suffers from sensitivity biases 
(Bolter et al. 2002). 

Potential problems encountered when enumerating microbes in soil 
include autofluorescence of soil matrix components, particularly in oil- 
contaminated soils, and occlusion of cells by soil particles, particularly 
clay-sized particles. In the latter case, methods have been developed to 
reduce interference by clays (Boenigk 2004) and confocal laser-scanning 
microscopy (CLSM) has been used to overcome problems of limited depth- 
of-focus in conventional microscopy. 

■ Equipment 

• Filter membranes (0.2 p.m pore size) for sterilizing reagents 

• Black polycarbonate filter membranes (0.2 p.m pore size, 25 mm diame- 
ter, e.g., Millipore; Millipore Corp., Billerica, MA, USA) 

• 25-mm filter holder unit consisting of a 15-mL glass reservoir and fritted 
glass base (wrapped and heat sterilized), clamp, and vacuum flask 

• Blunt-tipped filter forceps for handling filter membranes 



266 J. Foght, J. Aislabie 

• Vacuum pump with fine control 

• Glass microscope slides and coverslips, pre-cleaned 

• Epifluorescence microscope with appropriate filters 

■ Reagents 

• All diluents and reagents sterile and particle-free by filtration through 
0.2-|^m pore size membrane filters 

• Appropriate diluent for sample (Sect. 13.1) 

• Fluorescent stains appropriate to target cells: e.g., DAPI stock solution 
( 1 mg/mL) in deionized water, freshly diluted to a working concentration 
of 1 jig/mL in filtered deionized water, stains protected from light 

• Suitable wash solution: e.g., phosphate wash solution (PWS) containing 
10 mM KH 2 P0 4 , 0.85% NaCl and 5 mM MgCl 2 • 6H 2 

• Non-fluorescent immersion oil 



■ Sample Preparation 

Prepare suitable dilutions of soil sample (Sect. 13.1) in sterile, particle-free 
diluent. 



■ Procedure 

1. Prepare dilution series as required in filter-sterilized diluent. Vigorously 
mix sample for 5 min and allow suspension to stand for approx. 30 min 
to let larger soil particles settle out. If the sample will be kept longer 
than 30 min before counting, add a preservative (e.g., filter-sterilized 
formaldehyde to final concentration 3.7% or electron-microscopy-grade 
glutaraldehyde to final concentration 2.5%). 

2. Place black filter membrane in filter unit, add PWS (e.g., 4 ml) to column 
reservoir and known volume (e.g., 0.1 mL) of diluted soil suspension, 
avoiding settled soil particles. Perform subsequent steps under reduced 
lighting for light-sensitive stains like DAPI. 

3. Add required volume of stain (e.g., 1 mL DAPI working solution) to 
sample in column reservoir and stain in the dark for 7-10 min. 

4. Filter slowly through membrane under gentle vacuum. Rinse sides of 
column reservoir gently with diluent (two- to three-fold of initial volume) 
and allow filter to air dry. 



1 3 Enumeration of Soil Microorganisms 267 

5. Place a drop of immersion oil on a glass microscope slide, place the 
membrane filter on top, and cover with a coverslip. Follow with a drop 
of immersion oil and examine under an epifluorescence microscope at 
correct wavelength with appropriate filters. 

6. Count at least 20 fields of view (FOV) each containing 20-50 cells. Count 
randomly located FOV covering a wide area of the filter, avoiding its 
edges. 

7. Blanks consisting only of reagents should be performed at intervals, or at 
least at the beginning and end of sample enumeration. Blanks should be 
< 5% of the total cell densities in the samples and should be subtracted 
from sample counts before calculation of total numbers. 

■ Calculation 

Counts are calculated on the basis of wet mass of soil, corrected for back- 
ground, and usually expressed on the basis of dry mass of soil. 

- Cells/g soil wet mass = 

total no. of cells counted total stained area 1 

x x 



total no. of FOV area of FOV mass of soil on filter 

- Cells/g soil dry mass = 

(cells/g soil wet mass) x (dry-mass conversion factor) 

A specific example is given: 

- Area of FOV = 0.01 mm 2 

- Stained area of filter = nr 2 = 176.8 mm 2 

(diameter of the filter area covered by filtrate = 15 mm) 

- Total counts in 20 FOV for 0.1 mL of 10" 3 dilution = 929 

- Total counts in 20 FOV for reagent blanks = 40 

- Mass of soil on filter = 0.1 mL of 10~ 3 dilution = 10~ 4 g soil wet mass 

- Dry mass conversion factor (Sect. 13.1) = 1. 18 
Cells/g soil wet mass = 

(929 - 40) cells 176.8 mm 2 1 q 
x x - 7 9 x 10 

20 FOV 0.01mm 2 10" 4 g 

- Corrected count = 7.9 x 10 9 x 1. 18 = 9.3 x 10 9 cells/g soil dry mass 



268 J. Foght, J. Aislabie 

■ Notes and Points to Watch 

• An analysis of the sources of variation in the direct count method (Kirch- 
man et al. 1982) emphasizes the importance of enumerating replicate 
filters to reduce error. 

• Starving ("dwarf) cells and ultramicrobacteria (< 0.5 pm diameter) 
may not be not retained on the filter membrane or may not be detected 
by activity stains (Bolter et al. 2002). 

• At low cell densities it is difficult to achieve statistically valid counts, and 
efforts must be made to concentrate the sample if possible. 

• Hydrocarbon-contaminated samples may suffer from autofluorescence 
and poor disruption of aggregates. 

13.3 

Enumeration by Culture in Liquid Medium 

(Most Probable Number Technique) 

■ Introduction 

Objectives. The Most Probable Number (MPN) method uses statistics to 
infer the number of viable organisms in a sample that are able to grow or 
metabolize in a liquid medium under given incubation conditions. MPN 
tests can be carried out in large volumes in bottles or test tubes, or in 
microliter volumes in microtiter well plates, depending on the sample and 
the viability assay. 

Different media can be used to enumerate both generalist and specialist 
microbes in the soil. Total heterotrophs (generalists) can be enumerated in 
complex medium, although full-strength medium such as trypticase soy 
broth may not be suitable for enumerating microbes in nutrient-poor soils; 
for such samples tenth- strength medium maybe appropriate (Alef and Nan- 
nipieri 1995). The MPN method can be customized to differentiate among 
specialists by providing selective growth substrates. For example, mineral 
medium can be supplemented with filter-sterilized crude oil or refined 
product (e.g., diesel fuel) to enumerate "total hydrocarbon degraders" or 
amended with specific hydrocarbon substrates representing aliphatic and 
aromatic components (e.g., n-hexadecane and naphthalene, respectively). 
Liquid hydrocarbons can be added directly to broth whereas solid hydro- 
carbons can be provided as a fine suspension of crystals or dissolved in 
a non-metabolized water-immiscible carrier such as heptamethylnonane 
(Efroymson and Alexander 1991). Volatile hydrocarbons may be supplied 
in the vapor phase although this can be technically cumbersome. 



1 3 Enumeration of Soil Microorganisms 269 

Positive tubes maybe identified by various criteria, including: increased 
turbidity due to growth; emulsification of crude oil (e.g., "Sheen Screen," 
Brown and Braddock 1990); production of colored metabolites, particularly 
from some aromatic substrates (Stieber et al. 1994; Wrenn and Venosa 
1996); reduction of an iodonitrotetrazolium (INT) dye after incubation to 
indicate metabolism of substrates (Wrenn and Venosa 1996; Johnsen et al. 
2002); or evolution of 14 C0 2 from radiolabeled substrates (Carmichael and 
Pfaender 1997). It is important that both positive and negative controls be 
included with these tests. 

Principle. The microorganisms in a soil sample are serially diluted to ex- 
tinction, inoculated in replicate into a suitable medium, and incubated 
under appropriate conditions to yield a series of cultures that is scored 
according to pre-determined criteria. The combination of positive and 
negative cultures after incubation is evaluated by statistical methods to 
infer the MPN of viable cells in the undiluted sample. 

Theory. Culture-based enumeration methods such as MPN and plate count 
assay (Sect. 13.4) are biased because only a small proportion of environ- 
mental microbes has been cultured (Amann et al. 1995). With improved 
culture-based studies (e.g., Connon and Giovannoni 2002), the bias im- 
posed by growth-based methods will lessen, but it must be considered 
when interpreting results. The advantage to growth-based enumeration 
over molecular methods is that the former is technically simpler, usually 
easy to interpret, and can yield isolates for further investigation. The ad- 
vantage over plate count methods is that MPN is suitable for particulate 
samples (such as soil dilutions) that would obscure plate counts at low 
dilutions, and can detect microbes that will not grow on solid medium or 
are a minor component of a mixed culture. The disadvantages of MPN are 
that it yields only a statistical estimate of the viable microbes present and 
it requires many tubes and manipulations compared with plate counts. 

Typically a decimal dilution series is prepared in suitable diluent and 
a fixed volume of each dilution is inoculated into medium in replicate 
cultures, usually in multiples of 3, 5, or 10. MPN tests can be conducted in 
tubes, vials, or bottles, generally containing 7-10 mL medium per test tube, 
or in microtiter plates with 200 yiL per well. After incubation the tubes are 
scored qualitatively for criteria such as growth, production of metabolites, 
or loss of substrate. 

The combination of positive and negative cultures is converted to the 
MPN and confidence intervals either by consulting standard probability 
tables (e.g., Eaton et al. 1995; Alef and Nannipieri 1995) or using an algo- 
rithm (Koch 1994). The method assumes that (1) the microorganisms have 
been distributed into the cultures such that the highest dilution positive 
tubes were inoculated with a single organism, (2) culture tubes inoculated 



270 J. Foght, J. Aislabie 

with as few as one viable microbe will produce a positive result, and (3) the 
microbes have not been injured or rendered non-viable during sample 
handling. 

■ Equipment 

• Pipettes 

• Sterile test tubes or microtiter plates 

• Vortex mixer for mixing inoculum into medium (optional) 

• Incubation chamber with suitable temperature control and headspace 
(e.g., for anaerobes) 

• Microtiter plate reader for measuring color changes or optical density 
(optional) 

• Solvent-resistant filters (e.g., Millex-FG, Millipore Corp.) for filter ster- 
ilizing hydrocarbon solutions (optional) 

■ Reagents 

• Appropriate diluent for sample (Sect. 13.1) 

• Sterile liquid or semi-solid medium suitable for growth of target organ- 
ism^). For enumeration of generalists, standard or dilute liquid media 
(Alef and Nannipieri 1995, Atlas 1995) are appropriate; for enumeration 
of specialists, a mineral salts medium amended with selective carbon 
sources such as hydrocarbons maybe used (Sect. 13.4). 

• Specialty chemicals, depending on criteria for positive cultures, such as 
radiolabeled substrates, endpoint reagents, carrier solvents, etc. 

• Filter-sterilized liquid hydrocarbons or stock solutions of solid hydro- 
carbons dissolved in ethanol or dimethylformamide, for use as selective 
carbon sources (optional) 

■ Sample Preparation 

Perform serial dilutions of a representative soil sample in appropriate dilu- 
ent (Sect. 13.1), to exceed the expected viable number of cells by one or two 
orders of magnitude. 

■ Procedure 

1. Dispense replicate volumes of growth medium into suitable receptacles 
(e.g., 10 mL in test tubes, 200 pi per well for microtiter plates). Prepare 



1 3 Enumeration of Soil Microorganisms 27 1 

replicates (typically 3, 5, or 10) for each sample dilution to be tested. 
Medium must contain complete nutrients for growth including carbon 
source, and may contain indicators such as dyes or radiolabeled sub- 
strates. 

2. Inoculate replicate tubes with fixed volume of diluted sample (e.g., 1.0 mL 
for tubes, 100 pL for microtiter wells) covering at least three decimal 
dilutions. 

3. Include negative controls (uninoculated medium) and positive controls 
(medium inoculated with a culture known to produce a positive result) 
for reference. 

4. Incubate 7-14 days or longer in the dark under suitable conditions, 
taking into account in situ conditions of temperature, 2 levels, etc. 

5. Score tubes at intervals for positive results. Continue to incubate until 
two successive readings give the same results. Positive indicators include 
turbidity (e.g., heterotrophs growing in complex medium), hydrocarbon 
emulsification, production of soluble or gaseous metabolic end products 
(e.g., 14 C0 2 evolution from radiolabeled substrates, methane, colored 
metabolites), and changes in indicators (pH indicators, redox dyes). 

6. Identify the highest dilution set with all tubes positive, and the next two 
higher dilution sets. Use the pattern of positive and negative tubes with 
standard probability tables (e.g., Alef and Nannipieri 1995, Eaton et al. 
1995) to calculate the MPN from the dilution factor of the middle set. 
When non-standard patterns are encountered, follow the recommended 
variations provided with the tables for calculating the MPN. 

■ Calculation 

Published tables of statistical probability (Alef and Nannipieri 1995; Eaton 
et al. 1995; tables are also available on several internet sites such as US Food 
& Drug Administration) are used to convert the pattern of positive and 
negative tubes into the MPN of viable microbes in the original sample. The 
dilution factor and volume of sample used to inoculate the tubes are used in 
calculation but the volume of growth medium used in the tubes is not taken 
into consideration. Sample volumes reported in standard MPN tables are 
designed for water samples and are usually expressed per 1 00 mL of sample; 
therefore, they must be corrected for the actual volume of inoculum used 
in the test. Values are calculated as the MPN ± 95% confidence intervals 
(provided with the tables) and expressed on the basis of soil dry mass by 
multiplying the MPN by the dry-mass correction factor (Sect. 13.1). 

The simple algorithm below (Eaton et al. 1995) can be used to calcu- 
late the MPN without consulting published tables but does not provide 



272 J. Foght, J. Aislabie 

confidence intervals. 
MPN/lOOmL 

number of positive tubes x 100 
^(mL sample in negative tubes) (mL sample in all tubes) 

■ Notes and Points to Watch 

• Match the incubation conditions to in situ conditions when feasible. 
For example, select an appropriate culture incubation temperature (in- 
cluding temperature of the diluent and medium when inoculating), pro- 
vide semi-solid medium for enumeration of microaerophiles, anaerobic 
medium and headspace for anaerobes, etc. 

• If aerobic tubes are sealed, ensure that there is adequate headspace to 
maintain aerobic conditions if extended incubation will be required. 

• To use a high proportion of sample to growth medium, increase the 
strength of the medium (e.g., use double strength medium for 100% 
(v/v) inoculum). 

• When using turbidity as the criterion for growth, be aware of the tur- 
bidity contributed by soil particles at low dilutions, and by particulate 
substrates (e.g., suspensions of polycyclic aromatic hydrocarbon crys- 
tals). 

• If providing low molecular mass hydrocarbons as a carbon source avoid 
toxicity to the inoculum by minimizing substrate volumes. 

• Multiple MPN tests can be performed on a soil sample to enumerate 
different specialist components of the soil microbiota. If generalist MPN 
tests or direct counts (Sect. 13.2) are also performed, the specialists can 
be expressed as a proportion of the total viable numbers present in the 
sample. 

• After incubation, the positive MPN tubes may be suitable to use as an 
inoculum for subsequent isolation of pure cultures. 

13.4 

Enumeration by Culture on Solid Medium 

(Plate Count Technique) 

■ Introduction 

Objectives. The plate count technique quantifies the viable microbes in 
a sample by counting the number of colonies that form on or in a solid 



1 3 Enumeration of Soil Microorganisms 273 

growth medium inoculated with dilutions of that sample. Each colony is 
assumed to have originated from a single propagule or "colony forming 
unit" (CFU), whether that be a bacterial cell, endospore, hyphal fragment, 
or spore. Non-selective growth medium may be used to cultivate gener- 
alists, or selective medium may be used to enumerate specialists such as 
hydrocarbon-degrading bacteria and fungi. Specific enumeration of acti- 
nomycetes, filamentous fungi, or yeasts usually requires specialized media 
to suppress unwanted soil microbes such as spreading or mucoid colonies 
that overgrow the slower-growing colonies on non-selective plates (Labeda 
1990). Alternatively, a differential assay can be applied after the colonies 
have grown, to distinguish those possessing specific metabolic capabilities 
(e.g., production of colored metabolites; Kiyohara et al. 1982). Plates can be 
incubated under different atmospheres to enumerate aerobes, anaerobes, 
or microaerophiles, or at different temperatures to cultivate psychrotoler- 
ant, mesophilic, or thermophilic microbes. Plate counts maybe performed 
using several media and incubation conditions to enumerate different sub- 
sets of the viable microbes in a soil sample. 

Principle. Dilutions of a soil sample, performed in suitable diluent, are 
inoculated in replicate onto solid medium for cultivation with or without 
selection for specific metabolic types. Plates containing 30-300 colonies are 
selected and the colonies counted so that the CFU can be calculated for the 
original soil sample using dilution factors and dry-mass correction factors. 

Theory. It has long been recognized that the plate count method under- 
estimates the actual number of living cells in the sample by one or more 
orders of magnitude (Skinner et al. 1952) because soil organisms may not 
be viable or are not cultivable under the conditions employed (Amann et 
al. 1995). The proportion of viable cells enumerated will depend on the soil 
and on the growth medium and incubation conditions. New strategies for 
enumerating previously uncultured microbes are being devised to alleviate 
the cultivation bias (e.g., Joseph et al. 2003; Stevenson et al. 2004), but selec- 
tivity will always be a disadvantage of the plate count method (or any other 
cultivation-based enumeration method) compared with direct counts or 
molecular methods. The advantages are that the plate count method is rel- 
atively rapid and inexpensive and yields well-separated colonies suitable 
for subsequent purification and characterization. 

The medium and incubation conditions used in the plate count method 
determine which metabolic types of microbes will be enumerated, but 
the primary assumption for all plate counts is that each colony arises 
from a single viable propagule, i.e., a colony forming unit (CFU). After 
incubation, colonies are counted only on those plates containing 30-300 
colonies, for statistically valid enumeration (Koch 1994). Variations on the 
basic method exist, including spread plates appropriate for aerobic bacteria 



Vapor 


Spray 


Overk 


V 






V 








V 


V 




V 


V 




V 


V 



274 J. Foght, J. Aislabie 

Table 13.1. Recommended methods for providing hydrocarbons on solid medium 

Substrate Suitable for: 

Spread 
Toluene, xylenes 
Naphthalene 
Phenanthrene 
Dibenzothiophene 
Pyrene 

Octane +J 

Hexadecane *J 

Jet fuel aJ 

Crude oil *J 

and yeast, and pour plates suitable for microbes that do not grow well on 
surfaces but form subsurface colonies at reduced oxygen tension. Pour 
plates also help reduce problems with spreading colonies. Several methods 
can be used to supply substrates to mineral salts agar, including overlayer 
plates, spray plates and vapor plates (Table 13.1). Plates are incubated in 
the dark under suitable conditions of temperature and aeration, often for 
extended periods of time (e.g., 2-3 months) to enumerate slow-growing 
species (Janssen et al. 2002). 

■ Equipment 

• Sterile bent glass rod spreaders ("hockey sticks") for inoculating plates 

• Flame and beaker of ethanol to surface-sterilize spreaders 

• Manual turntable for turning Petri plates while spreading inoculum 
(optional) 

• Incubators with suitable atmosphere (aerobic or anaerobic) and temper- 
ature setting 

• Water bath at 50 °C for pour plates or overlayer plates 

• Aerosol spray apparatus (e.g., Jet-Pak, Sherwin-Williams Co., Cleveland) 
for applying ether solution of substrate to agar surface, or sealed con- 
tainers for incubating vapor plates (optional) 

■ Reagents 

• Solid growth medium in Petri plates, sufficient to inoculate 3 or 5 plates 
per dilution. Generalist media for bacteria include Plate Count Agar, Nu- 
trient Agar and R2A agar (Difco; Becton, Dickinson & Co., Sparks, MD, 



1 3 Enumeration of Soil Microorganisms 275 

USA) among many others. General mycological media include Czapek- 
Dox Agar (Difco), Malt Extract Agar (Difco) and Mycobiotic ("Mycosel") 
Agar (Acumedia, Neogen; Lansing, MI, USA), usually containing antibi- 
otics (e.g., oxytetracycline at lOOmg/L and/or streptomycin at 30mg/L) 
to suppress bacterial growth. Selective agar for hydrocarbon degraders is 
usually a mineral medium such as Bushnell Haas (Difco; Atlas 1995) so- 
lidified with 1.5% (w/v) Purified Agar (Oxoid, Basingstoke, UK) or Agar 
Noble (Difco) or 0.8% (w/v) gellan gum (Gelrite; Serva, Heidelberg) and 
amended with a specific carbon source. 

• Overlayer medium is usually prepared with agarose or purified agar 
(Bogardt and Hemmingsen 1992). 

• An ether or acetone solution of hydrocarbon substrate is used for spray 
plates. 



■ Sample Preparation 

Dilute sample appropriately in suitable diluent (Sect. 13.1) to exceed ex- 
pected number by at least one order of magnitude. 



■ Procedure 

Spread Plates (Bacteria, Yeasts, or Filamentous Fungi) 

1. Pipette a fixed volume of inoculum (typically 0.1 mL or 1.0 mL) from 
a range of dilutions onto three or five replicate agar plates and spread 
evenly using sterile bent glass rod. When inoculum has been absorbed 
into agar, invert plates and place in plastic bag to maintain humidity. 

2. Incubate at suitable temperature (e.g., ca. 25 °C for temperate soils) and 
under appropriate atmosphere, noting the appearance of colonies on 
plates with 30-300 colonies. Continue incubating until the number of 
colonies is constant. This may take less than a week for fast-growing 
bacteria, or more than 2 months for slow-growers or plates incubated 
at low temperatures. For extended incubation periods, seal the edges of 
plates with laboratory film or tape to prevent drying. 

3. Count the colonies on replicate plates having 30-300 colonies, determine 
the mean and calculate the CFU per gram dry mass in the original sample 
using the dilution factor and dry-mass correction factor. 

4. To enumerate colonies that can grow on liquid hydrocarbons spread 
a small volume (e.g., 50 p.L per plate) onto the surface of mineral medium 
agar either before or after inoculation, leaving small droplets on the agar 
surface. For solid hydrocarbons, see spray plate method below. 



276 J. Foght, J. Aislabie 

Pour Plates (Bacteria or Yeasts) 

1. Prepare 20 mL aliquots of molten growth medium agar and bring to ca. 
50 °C in a water bath. 

2. Add 1.0 mL inoculum to agar, mix briefly, and immediately pour into an 
empty sterile Petri plate. Alternatively, add inoculum to an empty sterile 
Petri plate, pour molten agar on top and mix by rotating the plate on the 
bench top. Allow agar to solidify then incubate as for spread plates. 

3. Count both surface and subsurface colonies. 

Overlayer Plates (Bogardt and Hemmingsen 1992) 

1. Prepare Petri plates containing 20 mL mineral medium with or without 
carbon source, depending on whether the overlayer contains a carbon 
source. 

2. Prepare 5mL sterile molten 1.5% (w/v) agarose or purified agar con- 
taining a suspension of particulate, insoluble substrate at a nominal 
concentration sufficient to provide an opaque suspension of fine crys- 
tals. It may be necessary to dissolve the substrate in a small volume of 
ethanol before adding to the agarose. Bring to 50 °C in a water bath. 

3. Add inoculum to the molten agarose, mix briefly, and immediately pour 
onto the mineral medium base, tipping the plate to distribute the over- 
layer evenly. The overlayer should be somewhat opaque. Allow to solidify 
then incubate as for spread plates. Alternatively, carefully inoculate the 
surface of the overlay as for spread plates. 

4. Count colonies that have a surrounding halo of clearing or colored 
metabolites (Fig. 13.1). 



Spray Plates for Solid Hydrocarbons (Kiyohara et al. 1982) 

1. Prepare a solution of crystalline hydrocarbon (e.g., phenanthrene or 
dibenzothiophene) in either acetone or anhydrous ethyl ether. CAU- 
TION: Ethyl ether is highly flammable and explosive. All procedures 
must be carried out in a well-vented fume hood away from any sparks or 
flames. Protective clothing and gloves must be worn to prevent exposure 
to hydrocarbon mist and precautions, such as spraying inside a card- 
board box in the hood, should be taken to prevent contaminating the 
fume hood with potentially carcinogenic compounds. The concentration 
of the solution does not need to be precise; approx. 10 mg of hydrocarbon 
dissolved in 2 mL of solvent should be sufficient to cover one plate. 

2. Use an aerosol canister to deliver a fine spray of solution to the surface 
of an inoculated plate. The surface should become slightly opaque with 



13 Enumeration of Soil Microorganisms 



277 




*t*> ^i . 






Fig. 13.1. Colonies capable of degrading 
carbazole form "haloes" of clearing in an 
overlayer plate prepared with carbazole. 
(From Shotbolt- Brown et al. 1996 with per- 
mission) 



a thin, even layer of very fine crystals. Seal edges of plates with laboratory 
film and place in plastic bags to prevent cross-contamination by vapors. 

3. Incubate and score for appearance of colonies. If the substrate can serve as 
a carbon source (e.g., phenanthrene), use mineral medium agar lacking 
a carbon source and score for colonies that are surrounded by zones of 
clearing and are larger than those observed on a parallel control plate 
lacking spray. If the substrate does not serve as a carbon source but can 
be co-metabolized (e.g., dibenzothiophene), use a low- nutrient agar that 
provides a carbon source and score for production of colored metabolites 
and/or zones of clearing around the colonies. 



Vapor Plates for Volatile Hydrocarbons 

1. Inoculate mineral medium agar by the spread plate method. 

2. For volatile solid hydrocarbons such as naphthalene, add a few crystals 
(< 0.1 g) to the lid of each inverted Petri plate, seal the edges of the 
plate with laboratory film, and incubate in sealed plastic bags to prevent 
cross-contamination on vapors. 

3. Volatile liquid hydrocarbons such as xylenes or jet fuel can be supplied 
to individual plates by placing a few drops of hydrocarbon into a plastic 
pipette tip stuffed with glass wool and placed on the lid of an inoculated, 
inverted mineral salts agar plate. Seal and incubate as above. To supply 
vapor to several plates at once, place inoculated plates into a sealable 
container with a small beaker containing a "wick" of glass wool or folded 
filter paper, and add a small amount of hydrocarbon, just sufficient to 



278 J. Foght, J. Aislabie 

saturate the headspace for a day or so, to reduce the chance of toxicity. 
Replenish hydrocarbon as necessary. 

■ Calculation 

1. Count the number of colonies arising on the plates, or the number 
showing the desired phenotype. 

2. Calculate the mean value for replicates, correct for dilution and dry mass, 
and express as CFU or phenotype-positive colonies per gram dry mass 
of original soil. 

■ Notes and Points to Watch 

• When spreading inoculum on a plate, make sure that the glass rod is not 
too hot. Similarly, do not use pour plates or overlayer plates to enumerate 
psychrophiles. 

• Incubate plates at temperatures close to those in situ when practical. 

• When enumerating specialist populations (e.g., hydrocarbon degraders), 
positive and negative controls should be included. 

• Antibiotics should be prepared as filter-sterilized concentrated solutions 
and added to cooled molten agar immediately before dispensing the agar 
into plates. Protect plates from the light before use and during incubation. 

• The use of low- nutrient media for enumeration has been recommended 
by some researchers, as discussed in Sect. 13.3. 

• Davis et al. (2005) suggest that plates with a minimum often colonies per 
plate rather than 30-300 colonies should be used for enumeration. This 
reduces depression in viable counts due to over-crowding of colonies on 
plates leading to inhibition of some species by others, or alternatively 
the depletion of nutrients by fast growing colonies that prevents slow 
growers from reaching a countable size. 

• Problems may arise with long-term incubation of plates for enumeration 
of slow-growing bacteria and fungi, including drying of the plates, ap- 
pearance of "spreading" or mucoid bacterial colonies or fungal colonies 
that obscure other colonies. 

• Plate counts over- represent genera that sporulate profusely (e.g., Peni- 
cillium, Trichoderma spp., Streptomyces spp.), and under-estimate or 
exclude fastidious genera. 

• Do not use too much solvent solution for spray plates, as the solvent can 
injure the inoculum. 



1 3 Enumeration of Soil Microorganisms 279 

References 

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demic Press, London 

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Brown EJ, Braddock JF (1990) Sheen screen, a miniaturized most-probable-number method 
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3896 

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Connon SA and Giovannoni SJ (2002) High-throughput methods for culturing microor- 
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Creach V, Baudoux A-C, Betru G, Le Rousiz B (2003) Direct estimate of active bacteria: CTC 
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14 



Quantification of Soil Microbial Biomass 
by Fumigation-Extraction 

Rainer Georg Joergensen, Philip C. Brookes 



14.1 

General Introduction 

The soil microbial biomass responds much more quickly than most other 
soil fractions to changing environmental conditions, such as variations 
in substrate input (e.g., Powlson et al. 1987) or increases in heavy metal 
content (Brookes and McGrath 1984). Much research supports the orig- 
inal idea of Powlson and Jenkinson (1976) that the biomass is a much 
more sensitive indicator of changing soil conditions than, for example, 
the total soil organic matter content. Thus the biomass can serve as an 
"early warning" of such changes, long before they are detectable in other 
ways. Biomass measurements are certainly useful in studies of soil pro- 
tection. They have the advantage that they are relatively cheap and sim- 
ple as well as being rapid. There is now a considerable amount of lit- 
erature to show that these measurements are useful in determining ef- 
fects of stresses on the soil ecosystem. Measurements of the soil microbial 
biomass by the fumigation extraction method have been used to estimate 
the environmental effects of pesticides (Harden et al. 1993) and antibiotics 
(Castro et al. 2002). This method has been repeatedly used to monitor 
successfully the bioremediation process of fuel oil contaminated soil (Joer- 
gensen et al. 1 994a, 1 994b, 1995,1997; Plante and Voroney 1 998; Franco et al. 
2004). 

Linked parameters (e.g., biomass-specific respiration or biomass as 
a percentage of soil organic C) are also useful because they possess "internal 
controls" (see Barajas Aceves et al. 1999 for a discussion). This fact permits 
interpretation of measurements in the natural environment, where, unlike 
in controlled experiments, there may not be suitable non-contaminated soil 
(for example) to provide good "control" or "background" measurements 
(Brookes 1995). 



Rainer Georg Joergensen: Department of Soil Biology and Plant Nutrition, University of Kas- 
sel, Nordbahnhofstr. la, 37213 Witzenhausen, Germany, E-mail: joerge@wiz.uni-kassel.de 

Philip C. Brookes: Agriculture and Environment Division, Rothamsted Research, Harpen- 
den, Herts., AL5 2JQ, UK 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



282 R.G. Joergensen, RC. Brookes 

14.2 

Fumigation and Extraction 

■ Introduction 

Objectives. The biomass of a microbial community can be quantitatively 
determined by fumigation and extraction in a large variety of soils devel- 
oped under very different environmental conditions, especially in contam- 
inated and remediated soils. 

Principle. Soils are fumigated with chloroform, incubated for 24 h, and ex- 
tracted. Different components can then be measured in the extracts, using 
various methods (Sects. 14.3-14.4). Non-fumigated soil is also extracted to 
correct for non-biomass soil organic matter. 

Theory. Following chloroform fumigation of soil, there is an increase in 
the amount of various organic and inorganic components coming from 
the cells of soil microorganisms (Jenkinson and Powlson 1976). The mem- 
branes of living soil microorganisms are partially lysed by the fumigant 
chloroform. After a 24 h incubation period to allow autolysis, a large part 
of the soil microbial biomass can be extracted from fumigated soil. The 
amount additionally rendered extractable from killed microorganisms is 
proportional to the original microbial biomass. Organic C (Vance et al. 
1987), total N and NH 4 -N (Brookes et al. 1985), and ninhydrin-reactive N 
(Joergensen and Brookes 1990) can be measured in the same 0.5 M K 2 S0 4 
extract (Alef and Nannipieri 1995). Organic C (Joergensen 1995) and total 
S (Wu et al. 1993) can be measured after extraction with 0.01 M CaCl 2 and 
phosphate or total P after extraction with NaHC0 3 (Brookes et al. 1982). 

■ Equipment 

• Room, incubator, or water bath adjustable to 25 °C 

• Implosion-protected desiccator 

• Vacuum line (water pump or electric pump) 

• Horizontal or overhead shaker 

• Deep-freezer at -15 °C 

• Folded filter papers (e.g., Whatman 42 or Schleicher & Schuell 595 1/2) 

• Glass conical flasks (250 mL) 



14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 283 

■ Reagents 

• Ethanol-free chloroform (CHCI3) 

• Soda lime 

• 0.5MK 2 SO 4 

■ Sample Preparation 

Use field-moist, sieved (between < 2 and < 5 mm) soil. 

■ Procedure 

1. Divide a moist soil sample of 50 g into two subsamples of 25 g. 

2. Place the non-fumigated control samples in 250 mL conical flasks, extract 
immediately with 100 mL 0.5 M K 2 S0 4 (extractant-to-soil ratio of 4:1) 
for 30 min by oscillating shaking at 200 rpm (or 45 min overhead shaking 
at 40 rpm), filter through a folded filter paper. 

3. For the fumigation treatment, place 50-mL glass vials containing the 
moist soil into a desiccator containing wet tissue paper and a vial of 
soda lime, add a beaker containing 25 mL ethanol-free CHC1 3 and a few 
boiling chips and evacuate the desiccator until the CHC1 3 has boiled 
vigorously for 2 min. 

4. Incubate the desiccator in the dark at 25 °C for 24 h. After fumigation, 
remove CHC1 3 by repeated (six- fold) evacuation and extract with 0.5 M 
K 2 S0 4 as described above. 

5. Store 0.5 M K 2 S0 4 extracts at -15 °C prior to analysis of organic C, total 
N, or ninhydrin-reactive N. 

■ Notes and Points to Watch 

• The desiccator must be kept under vacuum for 24 h to ensure the presence 
of a CHCI3 atmosphere, which kills virtually all soil microorganisms. 

• Ethanol-free CHC1 3 must be used to measure microbial biomass C be- 
cause ethanol cannot be completely removed from the soil after fumiga- 
tion. Ethanol-stabilized CHC1 3 can be used if solely microbial biomass N 
or ninhydrin-reactive N will be measured (DeLuca and Keeney 1993). 

• The soil must be sieved only if homogeneous samples are required (Ocio 
and Brookes 1990). 



284 R.G. Joergensen, RC. Brookes 

• Soil mass can range from 200 mg (Daniel and Anderson 1992) to 200 g 
(Ocio and Brookes 1990). 

• Soil microbial biomass is extracted by 0.5 M K 2 S0 4 . The high potas- 
sium concentration flocculates the soil and prevents adsorption of NHj 
released by fumigation. The relatively high salt concentration also in- 
hibits decomposition of the microbial material extracted after fumiga- 
tion. However, if the extracts have to be stored for a long period, they 
must be frozen. 

• Upon thawing of frozen K 2 S0 4 soil extracts, a white precipitate of CaS0 4 
occurs in near-neutral or alkaline soils. However, this causes no analytical 
problems in either method and may be safely ignored (Joergensen and 
Olfs 1998). 

• Soil water content can fluctuate widely, but must be higher than 30% 
water-holding capacity (WHC). Microbial biomass C and biomass N of 
soils at 40-50% WHC have been found to be similar to those in saturated 
soils (Widmer et al. 1989; Mueller et al. 1992). 

• Problems arise for fumigation and extraction in very compressed soils 
that cannot be dispersed. 

• Young living root cells are also affected by CHC1 3 fumigation. Conse- 
quently, in soils containing large amounts of living roots, a pre-extraction 
procedure must be carried out (Mueller et al. 1992). 

• In substrates containing more than 20% organic matter, e.g., compost, 
the ratio extractant-to-soil should be increased to 25:1 or more (Joer- 
gensen et al. 1997). 

14.3 
Biomass C 

■ Introduction 

Objectives. Very low concentrations of organic C can be measured in 0.5 M 
K 2 S0 4 soil extracts of fumigated and non-fumigated soil samples for the 
quantitative determination of soil microbial biomass C. 

14.3.1 

Biomass C by Dichromate Oxidation 

Principle. Organic C in the extracts is oxidized by dichromate digestion. 
The amount of dichromate left is determined after redox titration by the 
change in color from violet to dark green. 



14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 285 

Theory. In the presence of a strong acid and dichromate, organic matter 
is oxidized and Cr(+VI) reduced to Cr(+III). The amount of dichromate 
left is back-titrated with an iron II ammonium sulfate complex solution 
(Kalembasa and Jenkinson 1 973 ) and the amount of C oxidized is calculated. 

■ Equipment 

• Condenser 

• 250-mL round-bottom flask 

• Burette 

■ Reagents 

• K 2 Cr 2 7 solution (66.7 mM = 0.4 N) 

• Digestion mixture: Mix two parts cone. H 2 S0 4 with one part cone. H 3 P0 4 
(v/v) 

• Indicator solution: 0.1% Aldrich (Milwaukee) ferroin solution (1,10- 
phenanthroline-iron II sulfate complex) 

• Titration solution: 40 mM iron II ammonium sulfate [(NH 4 )2Fe(S0 4 )2 x 
6H 2 0] solution dissolved in distilled water, acidified with 20 mL cone. 
H 2 S0 4 , and made up to 1 L with distilled water 

■ Sample Preparation 

Use soil extract prepared as described in Sect. 14.2. 

■ Procedure 

1. Add 2mL of K 2 Cr 2 7 solution and 15mL of the digestion mixture to 
8 mL soil extract in a 250-mL round-bottom flask. 

2. Reflux the mixture gently for 30min, allow to cool, and dilute with 
20-25 mL water, added through the condenser as a rinse. 

3. Back-titrate excess dichromate with titration solution after adding 5 
drops of indicator solution to the digested soil extract. 

■ Calculation 

1. Calculation of extractable organic C 

(HB-S)xN xExVDx (VK + SW) x 1000 



C (iig/g soil) = 



CBxVSx DM 

(14.1) 



286 R.G. Joergensen, RC. Brookes 

HB consumption of titration solution by the hot (refluxed) blank (mL) 

S consumption of titration solution by the sample (mL) 

N normality of the K 2 Cr 2 7 solution 

E 3; conversion of Cr(+VI) to Cr(+III) assuming all organic C on 
average as [C(0)] 

VD added volume of the K 2 Cr 2 7 solution (mL) 

VS added volume of the sample (mL) 

VK volume of K 2 S0 4 extractant (mL) 

CB consumption of titration solution by the cold (unrefluxed) blank 
(mL) 

SW total amount of water in the soil sample (mL) 

DM total mass of dry soil sample (g) 

2. Calculation of microbial biomass C 

Biomass C = E c /k^c (14.2) 

Eq (organic C extracted from fumigated soils) 
- (organic C extracted non-fumigated soils) 

k EC 0.38 (Vance et al. 1987) 

■ Notes and Points to Watch 

• Be careful when working with K 2 Cr 2 7 ! 

• It is impossible to measure organic C with K 2 Cr 2 7 in the presence of 
high chloride concentrations. 

14.3.2 

Biomass C by UV-Persulfate Oxidation 

Principle. After removal of inorganic C by acidification, organic C in the 
extracts is oxidized by UV light at 210-260 nm in the presence of K 2 S 2 8 
to C0 2 , which is measured using an infrared absorption detector. 

Theory. The part of the microbial biomass rendered extractable after 
CHC1 3 fumigation is easily decomposable. For this reason, it is completely 
oxidized to C0 2 by UV-light in the presence of K 2 S 2 8 . Infrared strongly 
absorbs C0 2 . 



14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 287 

■ Equipment 

• Automatic carbon analyzer with IR-detection (e.g., Dohrman DC 80, 
Tekmar-Dohrmann, Cincinnati) 

■ Reagents 

• Acidification buffer: 50 g sodium hexametaphosphate [((NaP0 4 ) 6 )n] dis- 
solved in 900 mL distilled water, acidified to pH 2 with cone. H 3 P0 4 and 
made up to 1 L 

• Oxidation reagent: 20 g K 2 S 2 8 dissolved in 900 mL distilled water, acid- 
ified to pH 2 with cone. H 3 P0 4 and made up to 1 L 

■ Sample Preparation 

Use soil extract prepared as described in Sect. 14.2. 

■ Procedure 

For the automated UV-persulfate oxidation method, mix 5 mL K 2 S0 4 soil 
extract with 5 mL acidification buffer. Any precipitate of CaS0 4 in the soil 
extracts is dissolved by this procedure. The oxidation reagent is automat- 
ically fed into the UV oxidation chamber, where the oxidation to C0 2 is 
activated by UV light. The resulting C0 2 is measured by IR absorption. The 
IR detectors of Dimatec (Essen, Germany), for example, use a wavelength 
of 4.45 yarn (80 nm width) with 3.95 yam as reference. 

■ Calculation 

1. Calculation of extractable organic C 

[(SxDS)-(BxDB)]x(VK + SW) 

C(ug/gsoil) = (14.3) 

r& b DM 

S C in sample extract (yig/mL) 

B C in blank extract (yig/mL) 

DS dilution of sample with the acidification buffer 

DB dilution of blank with the acidification buffer 

VK volume of K 2 S0 4 extractant (mL) 

SW total amount of water in the soil sample (mL) 

DM total mass of dry soil sample (g) 



288 R.G. Joergensen, RC. Brookes 

2. Calculation of microbial biomass C 

Biomass C = E c /k EC (14.4) 

Eq (organic C extracted from fumigated soils) 
- (organic C extracted non-fumigated soils) 

k EC 0.45 (Wu et al. 1990; Joergensen 1996a) 

■ Notes and Points to Watch 

• It is impossible to measure organic C with the UV-persulphate oxidation 
method in the presence of high chloride concentrations because chloride 
absorbs a large amount of energy in the UV range. 

14.3.3 

Biomass C by Oven Oxidation 

Principle. After removal of inorganic C by acidification, organic C in the 
extracts is oxidized at 850 °C in the presence of platinum catalyzer to C0 2 , 
which is measured using an infrared absorption detector. 

Theory. Easily decomposable material as the part of the soil microbial 
biomass extractable after CHC1 3 fumigation is completely oxidized at 
850 °C in the presence of platinum catalyzer. Infrared strongly absorbs 
C0 2 . The new auto analyzers with oven systems [Shimadzu 5050 (Shi- 
madzu, Kyoto), Dimatoc 100 (Dimatec, Essen), multi N/C 2100 S (Analytik 
Jena, Jena, Germany), Maihack Tocor 4, Tocor 200 (SICK, Dusseldorf)] 
use small sample volumes so that they are able to measure C in extracts 
containing large amounts of salts (Joergensen and Olfs 1998). 

■ Equipment 

Automatic carbon analyzer with oven systems (for example Shimadzu 5050, 
Dimatoc 100, Dimatoc 2000, Analytik Jena multi N/C 2100 S; Analytik Jena 
multi N/C 3100), see manuals for detailed description. 

■ Sample Preparation 

Use soil extract prepared as described in Sect. 14.2. 

■ Procedure 

Dilute the samples to fit the calibration line and acidify using a few drops 
ofHCl. 



14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 289 

■ Calculation 

See Sect. 14.3.1. 

14.4 
Biomass N 

■ Introduction 

Objectives. Low concentrations of ninhydrin-reactive N or total N can be 
measured in 0.5 M K 2 S0 4 soil extracts of fumigated and non-fumigated soil 
samples without or with a digestion or oxidation step for the quantitative 
determination of soil microbial biomass N. 

14.4.1 

Ninhydrin-Reactive Nitrogen 

Principle. The amount of ninhydrin-reactive compounds released from the 
microbial biomass during the CHC1 3 fumigation and extraction by 0.5 M 
K 2 S0 4 is closely correlated to the initial soil microbial biomass C and 
biomass N content (Joergensen and Brookes 1991). 

Theory. Ninhydrin forms a purple complex with molecules containing a- 
amino nitrogen and with NHj and other compounds with free a-amino 
groups such as amino acids, peptides, and proteins (Moore and Stein 1948). 
The presence of reduced ninhydrin (hydrindantin) is essential to obtain 
quantitative color development with NHJ . 

■ Equipment 

• Boiling water bath 

• Spectrophotometer 

■ Reagents 

• Lithium acetate buffer (4 M, pH 5.2): 408 g lithium acetate dihydrate (for 
amino acid analysis) dissolved in water (400 mL), adjusted to pH 5.2 with 
96% acetic acid, and finally made up to 1 L with water 

• Citric acid buffer: citric acid monohydrate (42 g) and NaOH (16 g) dis- 
solved in water (900 mL), adjusted to pH 5 with 10 M NaOH if required, 
then finally made up to 1 L with water 



290 R.G. Joergensen, RC. Brookes 

• Ninhydrin reagent: 2 g ninhydrin and 0.3 g hydrindantin dihydrate dis- 
solved (for amino acid analysis) in 75 mL dimethylsulfoxide (DMSO), 
25 mL of 4 M lithium acetate buffer then added (Moore 1968) 

• Ethanol/water mixture (1 + 1, v/v) 

• Standard solutions: 10 mM L-leucine prepared in 0.5 M K 2 S0 4 and di- 
luted within the range 0-1,000 ]iM 



■ Sample Preparation 

Use soil extract prepared as described in Sect. 14.2. 

■ Procedure 

1. Add 0.6 mL of standard solutions, K 2 S0 4 soil extracts or blank, and 
1.4 mL of citric acid buffer to 20 mL test tubes (Joergensen and Brookes 
1990). 

2. Add 1 mL of ninhydrin reagent slowly, mix thoroughly, and close with 
loose aluminum lids. 

3. Heat the test tubes for 25 min in a vigorously boiling water bath; any 
precipitate formed during the addition of the reagents then dissolves. 

4. After heating, add 4 mL of the ethanol-to-water mixture, mix the solu- 
tions thoroughly, and read the absorbance at 570 nm. 

■ Calculation 

1. Calculation of extracted ninhydrin-reactive N (N n i n ) 

xt / / -n (S-B)xNx(VK + SW) 

N nin (jig/g soil) = — (14.5) 

L x DM 

S absorbance of the sample 

B absorbance of the blank 

JV atomic mass of nitrogen (14) 

VK volume of K 2 S0 4 extractant (mL) 

SW total amount of water in the soil sample (mL) 

L millimolar absorbance coefficient of leucine 

DM total mass of dry soil sample (g) 



14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 291 

2. Calculation of microbial ninhydrin-reactive N 

Bnin - (N n in extracted from the fumigated soil) 

(14.6) 
_ (N n in extracted from the non-fumigated soil) 

3. Calculation of microbial biomass C 

Biomass C = B n i n x 22 

(for soils with a pH( H2 o) above 5.0; Joergensen 1996b) 

Biomass C = B n i n x 35 

(for soils with a pH( H2 o) of or below 5.0; Joergensen 1996b) 

■ Notes and Points to Watch 

• A reflux digestion is not required for ninhydrin N. This makes it very 
suitable for situations with minimal laboratory facilities. 

• In both biomass C and N measurements the fraction coming from the 
biomass is determined following subtraction of an appropriate "control." 
With biomass C this value is often half of the total, while with biomass 
ninhydrin N it is commonly about 10% or less. This causes considerably 
less error in its determination. 

• At 100 °C the reaction with free amino groups of proteins and amino 
acids is essentially complete within 1 5 min (e.g., leucine reaches the max- 
imum optical density after approximately 5 min). However the reaction 
of hydrindantin with NHj requires 25 min. 

• The ratio between the volume of the sample and that of citric acid should 
not be closer than 0.75:1.75 to avoid the formation of a precipitate after 
the addition of the ninhydrin reagent. 

• The most common solvent in the ninhydrin method is 2-methoxyethanol 
(Amato and Ladd 1998). However, because it is an ether it tends to 
form peroxides that destroy ninhydrin and hydrindantin. Dimethylsul- 
foxide (DMSO) is peroxide free, has lower toxicity and a higher boil- 
ing point (189 °C), and gives a more stable color development than 
2-methoxyethanol. 

• The ninhydrin method proposed by Amato and Ladd (1988) for 2M 
KC1 extracts does not require the use of citric acid buffer. The optimum 
reagent-to-sample ratio is 1:2. 



292 R.G. Joergensen, RC. Brookes 

14.4.2 

Total Nitrogen 

Principle. Total nitrogen is measured under strong acidic conditions by 
Kjeldahl digestion. Ammonium can be measured by distillation (see 
Chapt. 16). 

Theory. Ammonium is released from amines, peptides and amino acids in 
0.5 M K 2 S0 4 soil extracts of fumigated and non-fumigated soil samples. Ni- 
trate is additionally reduced to ammonium under strong acidic conditions 
in the presence of KCr(S0 4 ) 2 , Zn powder, and CuS0 4 as reducing agents. 

■ Equipment 

• Digestion block 

• Steam distillation apparatus 

• Burette or autotitrator 

■ Reagents 

• Reducing agent: 50 g of chromium(III) potassium sulfate dodecahydrate 
(KCr(S0 4 ) 2 x 12H 2 0) dissolved in approx. 700 mLdeionized water, and 
after adding 200 mL cone. H 2 S0 4 , cooled and diluted to 1,000 mL 

• Zn powder 

• CuS0 4 solution (0.1 9 M) 

• Cone. H 2 S0 4 

• lOMNaOH 

• 2% H3BO3 

• IO11MHCI 

■ Sample Preparation 

Use soil extract prepared as described in Sect. 14.2. 

■ Procedure 

1. Add 10 mL of the reducing agent and approx. 300 mg Zn powder to 30 mL 
of the K 2 S0 4 soil extract and leave for at least 2 h at room temperature. 

2. Add 0.6 mL of CuS0 4 solution, 8 mL of cone. H 2 S0 4 , heat gently for 2 h 
until all the water has disappeared, and then heat for 3 h at the maximum 
temperature. 



14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 293 

3. Allow the digest to cool before distillation with 40 mL 10 M NaOH. The 
evolved NH 3 is adsorbed in 2% H3BO3. 

4. Titrate the resulting solution with 10 pM HC1 to pH 4.8. 



■ Calculation 

1. Calculation of extractable total N 

1X (S-B)xMxNx(VK + SW) 

N pg/g soil = 14.7 

^ b b Ax DM 

S HC1 consumed by sample extract (pL) 

B HC1 consumed by blank extract (pL) 

M molarity of HC1 

N molecular mass of nitrogen (14) 

VK volume of K 2 S0 4 extractant (mL) 

SW total amount of water in the soil sample (mL) 

A sample aliquot (mL) 

DM total mass of dry soil sample (g) 

2. Calculation of microbial biomass N 

Biomass N = En/^en (14.8) 

£ N (total N extracted from fumigated soils) 
- (total N extracted non-fumigated soils) 

fc EN 0.54 (Brookes et al. 1985; Joergensen and Mueller 1996) 

■ Notes and Points to Watch 

• A method is available in which the extracted total N is oxidized to NO3 , 
which is then determined colorimetrically (Cabrera and Beare 1993). 

• If losses of NO3 occur during the fumigation period, they can be cor- 
rected by considering the difference between the NO3 extracted initially 
and the NO3 extracted at the end of the fumigation period (Brookes et al. 
1985). 

• If (non-fumigated) soil samples contain large amounts of NO3 or NHJ 
in the soil solution, a pre-extraction step should be carried out (Wid- 
mer et al. 1989; Mueller et al. 1992; Joergensen et al. 1995). 



294 R.G. Joergensen, P.C. Brookes 

References 

Alef K, Nannipieri P (1995) Methods in Applied Soil Microbiology and Biochemistry. Aca- 
demic Press, London 
Amato M, Ladd JN (1988) Assay for microbial biomass based on ninhydrin-reactive nitrogen 

in extracts of fumigated soils. Soil Biol Biochem 20:107-114 
Barajas Aceves M, Grace C, Ansorena J, Dendooven L, Brookes PC (1999) Soil microbial 

biomass and organic C in a gradient of zinc concentrations around a spoil tip mine. Soil 

Biol Biochem 31:867-876 
Brookes PC (1995) The use of microbial parameters in monitoring soil pollution by heavy 

metals. Biol Fertil Soils 19:269-279 
Brookes PC, Landman A, Pruden G, Jenkinson DS (1985) Chloroform fumigation and the 

release of soil nitrogen: A rapid direct extraction method for measuring microbial 

biomass nitrogen in soil. Soil Biol Biochem 17:837-842 
Brookes PC, McGrath SP (1984) The effects of metal toxicity on the soil microbial biomass. 

J Soil Sci 35:341-346 
Cabrera ML, Beare MH (1993) Alkaline persulfate oxidation for determining total nitrogen 

in microbial biomass extracts. Soil Sci Soc Am J 57:1007-1012 
Castro J, Sanchez-Brunete C, Rodriguez JA, Tadeo JL (2002) Persistence of chlorpyrifos and 

endosulfan in soil. Fres Environ Bull 11:578-582 
Daniel O, Anderson JM (1992) Microbial biomass and activity in contrasting soil materials 

after passage through the gut of the earthworm Lumbricus rubellus Hoffmeister. Soil 

Biol Biochem 24:465-470 
DeLuca TH, Keeney DR (1993) Ethanol-stabilized chloroform as fumigant for estimating 

microbial biomass by reaction with ninhydrin. Soil Biol Biochem 25:1297-1298 
Franco I, Contin M, Bragato G, De Nobili M (2004) Microbiological resilience of soils 

contaminated with crude oil. Geoderma 121:17-30 
Harden T, Joergensen RG, Meyer B, Wolters V (1993) Mineralization of straw and formation 

of soil microbial biomass in a soil treated with simazine and dinoterb. Soil Biol Biochem 

25:1273-1276 
Jenkinson DS, Powlson DS (1976) The effects of biocidal treatments on metabolism in soil - 

I. Fumigation with chloroform. Soil Biol Biochem 8:167-177 
Joergensen RG ( 1 995 ) The fumigation-extraction method to estimate soil microbial biomass: 

Extraction with 0.01 m CaCl 2 . Agribiol Res 48:319-324 
Joergensen RG (1996a) The fumigation-extraction method to estimate soil microbial 

biomass: Calibration of the k^c value. Soil Biol Biochem 28:25-31 
Joergensen RG (1996b) Quantification of the microbial biomass by determining ninhydrin- 
reactive N. Soil Biol Biochem 28:301-306 
Joergensen RG, Brookes PC (1990) Ninhydrin-reactive nitrogen measurements of microbial 

biomass in 0.5 M K2SO4 soil extracts. Soil Biol Biochem 22:1023-1027 
Joergensen RG, Figge RM, Kupsch L ( 1 997) Microbial decomposition of fuel oil after compost 

addition to soil. Z Pflanzenernahr Bodenk 160:21-24 
Joergensen RG, Mueller T (1996) The fumigation-extraction method to estimate soil micro- 
bial biomass: Calibration of the /cen value. Soil Biol Biochem 28:33-37 
Joergensen RG, Olfs HW (1998) The variability between different analytical procedures and 

laboratories for measuring soil microbial biomass C and biomass N by the fumigation 

extraction method. Z Pflanzenernahr Bodenk 161:51-58 
Joergensen, RG, Schmaedeke F, Windhorst K, Meyer B (1994a) Biomasse und Aktivitat 

von Mikroorganismen eines mineralolkontaminierten Bodens. In: Alef K, Fiedler H, 

Hutzinger O (eds) Band 6: Bodenkontamination, Bodensanierung, Bodeninformation- 

ssysteme. Eco-Informa'94, Umweltbundesamt/Wien, pp 225-236 



14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 295 

Joergensen RG, Schmaedeke F, Windhorst K, Meyer B ( 1 994b) Die Messung der mikrobiellen 

Biomasse wahrend der Sanierung eines mit Dieselol kontaminierten Bodens. VDLUFA- 

Schriftenr 38:557-560 
Joergensen RG, Schmaedeke F, Windhorst K, Meyer B (1995) Biomass and activity of mi- 
croorganisms in a fuel oil contaminated soil. Soil Biol Biochem 27:1137-1143 
Kalembasa SJ, Jenkinson DS (1973) A comparative study of titrimetric and gravimetric 

methods for the determination of organic carbon in soil. J Sci Food Agric 24:1085-1090 
Moore S (1968) Amino acid analysis: Aqueous dimethyl sulfoxide as solvent for the ninhydrin 

reaction. J Biol Chem 243:6281-6283 
Moore S, Stein WH (1948) Photometric ninhydrin method for use in the chromatography 

of amino acids. J Biol Chem 176:367-388 
Mueller T, Joergensen RG, Meyer B (1992) Estimation of soil microbial biomass C in the 

presence of living roots by fumigation-extraction. Soil Biol Biochem 24:179-181 
Ocio JA, Brookes PC (1990) Soil microbial biomass measurements in sieved and unsieved 

soils. Soil Biol Biochem 22:999-1000 
Plante AF, Voroney RP (1998) Decomposition of land applied oily food waste and associated 

changes in soil aggregate stability. J Environm Qual 27:395-402 
Powlson DS, Brookes PC, Christensen BT (1987) Measurement of soil microbial biomass 

provides an early indication of changes in total soil organic matter due to straw incor- 
poration. Soil Biol Biochem 19:159-164 
Powlson DS, Jenkinson DS (1976) The effects of biocidal treatments on metabolism in 

soil. II gamma irradiation, autoclaving, air-drying and fumigation. Soil Biol Biochem 

8:179-188 
Vance ED, Brookes PC, Jenkinson DS (1987) An extraction method for measuring soil 

microbial C. Soil Biol Biochem 19:703-708 
Widmer P, Brookes PC, Parry LC (1989) Microbial biomass nitrogen measurements in soils 

containing large amounts of inorganic nitrogen. Soil Biol Biochem 21:865-867 
Wu J, Joergensen RG, Pommerening B, Chaussod R, Brookes PC (1990) Measurement of soil 

microbial biomass C - an automated procedure. Soil Biol Biochem 22:1167-1169 
Wu J, O'Donnell AG, Syers JK (1993) Microbial growth and sulphur immobilization following 

the incorporation of plant residues into soil. Soil Biol Biochem 25:1567-1573 



15 



Determination of Adenylates 
and Adenylate Energy Charge 

Rainer Georg Joergensen, Markus Raubuch 



■ Introduction 

Objectives. The determination of adenosine-triphosphate (ATP) extracted 
from soil was introduced a long time ago as an estimate of the soil microbial 
biomass (Oades and Jenkinson 1979). After a conditioning pre-incubation, 
close linear relationships exist between ATP and microbial biomass C de- 
termined either by the fumigation incubation technique (Jenkinson 1988) 
or by the fumigation extraction method (Chapt. 14; Contin et al. 2001; Dy- 
ckmans et al. 2003). A similar close linear relationship exists also between 
microbial biomass C and the sum of all three adenylates AMP, ADP, and 
ATP (Dyckmans et al. 2003). The determination of adenylates is the quickest 
way of estimating microbial biomass, because 24-h incubation periods or 
manipulations such as substrate addition are not required as in the fumiga- 
tion extraction or the substrate induced respiration methods, respectively. 
The measurement of adenylates by high-performance liquid chromatog- 
raphy (HPLC) has been repeatedly used to monitor the effects of heavy 
metal contamination (Chander et al. 2001) and salinization (Sardinha et al. 
2003), but no information is available regarding fuel oil contaminated soil. 
However, enzymatic ATP has been successfully used to monitor microbial 
activity during fuel oil decomposition, although some quenching of the 
bioluminescence by fuel oil residues occurred (Wen et al. 2003). 

An important index for the energetic state of the soil microbial commu- 
nity is the adenylate energy charge (AEC), which was defined by Atkinson 
and Walton (1967) as follows: 

(ATP + 0.5 x ADP) /(ATP + ADP + AMP) 

High AEC values (> 0. 7) have frequently been described in soils (Brookes 
et al. 1987; Brookes 1995; Chander et al. 2001; Dyckmans et al. 2003). 
Low AEC values have been demonstrated under drought stress conditions 
(Raubuch et al. 2002), but also in Cu contaminated soils (Chander et al. 
2001) and in acidic saline soils (Sardinha et al. 2003). 



Rainer Georg Joergensen, Markus Raubuch: Department of Soil Biology and Plant Nu- 
trition, University of Kassel, Nordbahnhofstr. la, 37213 Witzenhausen, Germany, E-mail: 
joerge@wiz.uni-kassel.de 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



298 R.G. Joergensen, M. Raubuch 

Principle. Soil adenylates (AMP + ADP + ATP) are extracted with dimethyl- 
sulfoxide (DMSO) under strong alkaline conditions in combination with 
an ethylene-diamine-tetraacetic acid (EDTA)-containing phosphate buffer. 
DMSO destroys microbial cells, the phosphate buffer completely prevents 
the adsorption of adenylates under the strong alkaline conditions, and 
EDTA promotes the irreversible inactivation of ATP-converting enzymes. 

Theory. ATP is rapidly destroyed outside living cells and can be used as 
an estimate for the soil microbial biomass assuming a constant ATP-to- 
microbial biomass ratio, which is fairly true in the absence of living plant 
roots and after a conditioning pre-incubation (Jenkinson 1988). The ATP- 
to-microbial biomass C ratio is affected by drought (Raubuch et al. 2002), 
temperature (Joergensen and Raubuch 2003), and N limitation (Joergensen 
and Raubuch 2002). However, the main problems in measuring ATP in soils 
are (1) the enzymatic breakdown of ATP after cell death and (2) adsorp- 
tion of ATP to clay minerals during extraction (Martens 2001). The alka- 
line DMSO-EDTA-phosphate-buffer extractant solved nearly all method- 
ological problems reported earlier (Bai et al. 1988; Martens 1992). This 
is especially true in combination with HPLC analysis after derivatization 
with chloroacetaldehyde to form the fluorescent l-N 6 -etheno-derivatives 
(^-adenylates), which are highly selective for fluorometric determination 
(Bai et al. 1989; Dyckmans and Raubuch 1997). 

■ Equipment 

• Multipoint magnetic stirrer 

• Ultrasonic bath 

• Evacuation units and filters (0.45-^m cellulose nitrate membrane filters) 

• Heating water bath 

• Test tube stirrer 

• Glassware: 100-mL glass beaker (tall form), 20-mL test tubes 

• Pipettes 

• HPLC equipment: automatic injector, isocratic precision pump, column 
oven, solvent delivery system, fluorescence detector and recording unit 

• Analytical column (250 x 4.6 mm; 5 p.m ODS Hypersil, Thermo Electron 
Corp., Waltham, MA, USA) with guard column (10 x 4.0 mm, 5 pm ODS 
Hypersil) 



15 Determination of Adenylates and Adenylate Energy Charge 299 

■ Reagents 

• DMSO 

• Extraction buffer: 20 mM EDTA dissolved in 10 mM Na 3 P0 4 x 12H 2 
containing 0.1 M KOH at pH 12 

• Tris buffer: 2 mM EDTA dissolved in 10 mM ammonium acetate/20 mM 
Tr is (hydroxy methyl) -aminome thane, adjusted to pH 7.75 with acetic 
acid (store at 4°C) 

• Adenylate releasing reagent: 0.05 mL benzalkonium chloride solution 
(ca. 50% in water, Fluka, Fluka AG, Buchs, Switzerland, purum grade) 
added to 49.95 mL Tris buffer (store at 4 °C) 

• 0.1MKH 2 PO 4 

• Chloroacetaldehyde 

• TBAHS buffer: 50 mM ammonium acetate, 1 mM EDTA, 0.4 mm tetra- 
ft-butylammonium hydrogen sulfate (TBAHS, LiChropur, Merck KGaA, 
Darmstadt, Germany) 

• Mobile phase for HPLC: TBAHS buffer mixed with methanol at a ratio 
of89.5tol0.5(v/v) 

• Calibration stock solution I (lOOiig/mL): 14.35 mg AMP-Na 2 x 6H 2 0, 
11.59mgADP-K 2 x 2H 2 0, or 11.90 mgATP-Na 2 x 3H 2 0; each dissolved 
in 100 mL extraction buffer (store at 4 °C) 

• Calibration stock solution II (1 p.g/mL): 1/100 dilution of stock solution I 
(store at 4 °C) 

• Working standard solutions: a set of four standards each containing 2, 4, 
6, 8 ng of AMP, ADP, ATP, respectively, prepared by mixing 100-400 \xL 
stock solution II with 0.2 mL chloroacetaldehyde and adding 0.01 M 
Na 2 HP0 4 x 2H 2 to give a final volume of 10 mL, heated for 3min at 
85 °C, and cooled in an ice bath (store at 4 °C for maximum 7 days) 

■ Sample Preparation 

Use moist sample equivalent to 1-5 g oven-dry soil, sieved (< 2 mm). The 
experimental design reflects the fact that adenylate content responds to 
actual conditions, is influenced by mechanical disturbance, water content, 
and temperature. 



300 R.G. Joergensen, M. Raubuch 

■ Procedure 

1. Weigh moist soil equivalent to 1-5 g oven-dry soil into a 100-mL glass 
beaker (tall form). 

2. Add 4 mL DMSO and stir for 2 min on a magnetic stirrer using a mag- 
netic stirring bar. 

3. Add 16 mL extraction buffer and stir again for 2 min. 

4. Sonify for 2 min in an ultrasonic bath. 

5. Mix an aliquot of 0.5 mL of soil suspension with 0.5 mL of adenylate 
releasing reagent in a 20 mL test tube, mix using a test tube stirrer, and 
sonify for another 5 s. 

6. Pass the suspension through a membrane filter (0.45 p.m) and wash the 
soil residue twice with 1 mL 0.1 M KH 2 P0 4 . 

7. Add 0.2 mL chloroacetaldehyde and make up to a final volume of 5 mL 
by addition of 0.1 M KH 2 P0 4 . 

8. Incubate in a water bath for 30 min at 85 °C to yield the fluorescent 
l-N 6 -etheno-derivatives and cool afterward in an ice bath. 

9. Store at 4 °C for a maximum 7 days before HPLC measurements. 

10. Adjust the column oven to 27 °C. 

11. Run HPLC with the mobile phase at 2 mL/min for 3 h for equilibration 
of the column. 

12. Use a sample loop of 200 p.L. 

13. Fluorometric emission is measured at 410 nm with 280 nm as excitation 
wavelength. 

14. Clean the HPLC after measurement for 30 min at 1 mL/min with a meth- 
anol/water (50:50 v/v) solution. 

15. Treat calibration standards like soil extractants to prepare calibration 
curves. 

16. Standard solutions correspond to concentrations 2ng, 4ng, 6ng, 8ng 
of AMP, ADP and ATP in 200 ]iL y respectively. 

17. There is a linear relationship in adenylate content and signal response 
up to 8 ng of each adenylate. The adenylates are detected on the chro- 
matogram in the order AMP, ADP, and ATP. 



15 Determination of Adenylates and Adenylate Energy Charge 301 

■ Calculation 

1. Identify AMP, ADP and ATP by retention time according to the retention 
time of the standards. 

2. Calculate nanograms from areas and linear equation of standards. 

3. Take dilution into account (analogous for ADP and AMP). 

H x (E + SW) x I 



ATP (ng/gsoil) = 



A xDM 



H ATP in 200 p.L injection volume (ng) 

E extractant (4 mL DMSO + 16 mL extraction buffer; mL) 

J 25; conversion factor of the injection volume (200 p.L from 5 mL) 

SW total amount of water in the soil sample (mL) 

A aliquot (0.5 mL) 

DM total mass of dry soil sample (g) 

4. Molecular masses for conversion into nmol: 
AMP = 347.2 g, ADP = 427.2 g, ATP = 507.2 g 

5. Total adenylate content (nmol/g soil) = AMP + ADP + ATP 

6. Adenylate Energy Charge (AEC) 

= (ATP + 0.5 ADP)/(AMP + ADP + ATP) 



Notes and Points to Watch 

The mobile phase must be degassed in advance. Oxygen disturbs the 
measurement, especially of ATP. 

The retention time must be checked before the first measurement with 
a standard mixture of AMP, ADP, and ATP standards, but do not use 
a standard mixture of AMP, ADP and ATP for calibration. ATP contains 
impurities of AMP and ADP, ADP contains impurities of AMP. 

The column temperature should be constant at 27 °C. The separation of 
£-AMP, £-ADP and £-ATP from extracted impurities is improved at 27 °C. 
Changing temperatures causes shifts in the retention times. 



302 R.G. Joergensen, M. Raubuch 

References 

Atkinson DE, Walton GM (1967) Adenosine triphosphate conservation in metabolic regu- 
lation. Rat liver cleavage enzyme. J Biol Chem 242:3239-3241 

Bai QY, Zelles L, Scheunert I, Korte F (1988) A simple procedure for the determination of 
adenosine triphosphate in soils. Chemosphere 17:2461-2470 

Bai QY, Zelles L, Scheunert I, Korte F (1989) Determination of adenine nucleotides in soil by 
ion-paired reverse-phase high-performance liquid chromatography. J Microbiol Meth 
9:345-351 

Brookes PC (1995) Estimation of the adenylate energy charge in soils. In: Alef K, Nannipieri P 
(eds) Methods in Applied Soil Microbiology and Biochemistry. Academic Press, London, 
pp 204-213 

Brookes PC, Newcombe AD, Jenkinson DS (1987) Adenylate energy charge measurements 
in soil. Soil Biol Biochem 19:211-217 

Chander KC, Dyckmans J, Joergensen RG, Meyer B, Raubuch M (2001) Different sources of 
heavy metals and their long-term effects on soil microbial properties. Biol Fertil Soils 
34:241-247 

Contin M, Todd A, Brookes PC (2001) The ATP concentration in the soil microbial biomass. 
Soil Biol Biochem 33:701-704 

Dyckmans J, Chander K, Joergensen RG, Priess J, Raubuch M, Sehy U (2003) Adenylates as an 
estimate of microbial biomass C in different soil groups. Soil Biol Biochem 35:1485-1491 

Dyckmans J, Raubuch M (1997) A modification of a method to determine adenosine nu- 
cleotides in forest organic layers and mineral soils by ion-paired reversed-phase high- 
performance liquid chromatography. J Microbiol Meth 30:13-20 

Jenkinson DS (1988) The determination of microbial biomass carbon and nitrogen in soil. 
In: Wilson JR (ed) Advances in nitrogen cycling in agricultural ecosystems. CABI, 
Wallingford, pp 368-386 

Joergensen RG, Raubuch M (2002) Adenylate energy charge of a glucose-treated soil without 
adding a nitrogen source. Soil Biol Biochem 34:1317-1324 

Joergensen RG, Raubuch M (2003) Adenylate in the soil microbial biomass at different 
temperatures. Soil Biol Biochem 35:1063-1069 

Martens R (1992) A comparison of soil adenine nucleotide measurements by HPLC and 
enzymatic analysis. Soil Biol Biochem 24:639-645 

Martens R (200 1 ) Estimation of ATP in soil: extraction methods and calculation of extraction 
efficiency. Soil Biol Biochem 33:973-982 

Oades JM, Jenkinson DS (1979) Adenosine triphosphate content of the soil microbial 
biomass. Soil Biol Biochem 11:193-199 

Raubuch M, Dyckmans J, Joergensen RG, Kreutzfeldt M (2002) Relation between respi- 
ration, ATP content and adenylate energy charge (AEC) after incubation at different 
temperatures and after drying and rewetting. J Plant Nutr Soil Sci 165:435-440 

Sardinha M, Miiller T, Schmeisky H, Joergensen RG (2003) Microbial performance in a tem- 
perate floodplain soil along a salinity gradient. Appl Soil Ecol 23:237-244 

Wen G, Voroney RP, McGonigle TP, Inanaga S (2003) Can ATP be measured in soils treated 
with industrial oily waste? J Plant Nutr Soil Sci 166:724-730 



16 



Determination 

of Aerobic N-Mineralization 

Rainer Georg Joergensen 



■ Introduction 

Objectives. N mineralization is the transformation of organic N into inor- 
ganic N components (Beck 1983). It is thus an important biological process 
of the nitrogen cycle in ecosystems, reflecting the ability of a soil to provide 
available nitrogen to plants (Alef 1995). In terrestrial ecosystems, especially 
in arable crop production, N is often the most limiting nutrient for plant 
growth. The nitrogen availability to soil microorganisms often limits the 
decomposition of fuel oil in contaminated soils (Joergensen et al. 1995). N 
mineralization mainly depends on temperature, moisture, aeration, type 
of organic N, and pH. NHJ is subject to fixation by clays. NO3 can be lost 
through denitrification and leaching (Alef 1995). 

Principle. A soil is incubated aerobically after removal of plant debris for 
two periods. The soil is extracted with 2 M KC1 before and after each of the 
two incubation periods. In the soil extracts, NHj, NO3, and, if necessary, 
NO2 are measured. 

Theory. N mineralization is the catabolic use of N- containing organic com- 
ponents, e.g., amino acids, amino sugars, amines, and nucleic acids derived 
from plant and animal debris as well as from soil organic matter to meet 
the energy demand of the soil microbial biomass. The N mineralization 
process can be divided into two steps (Alef 1995): (1) the first step is am- 
monification, which is the breakdown of organic NH 2 groups to NHJ. 
Except for the hydrolysis of urea by extracellular urease, ammonification 
is carried out by proteases bound to cell membranes of all heterotrophic 
microorganisms in soil, i.e. more than 95% of the soil microbial com- 
munity. (2) The second step is nitrification, which is carried out by het- 
erotrophic fungi in acidic soils or obligatory aerobic chemoautotrophic 
bacteria [e.g., Nitrosomonas: NHJ —> NO2 (GO = -273.9 kj/mol) andM- 
trobacter NO2 —> NO3 (GO = -76.7 kj/mol) under neutral and slightly 
alkaline soil conditions. Nitrification is inhibited by fuel oil contamination 
in contrast to ammonification (Joergensen et al. 1995). In compacted soils, 



Rainer Georg Joergensen: Department of Soil Biology and Plant Nutrition, University of Kas- 
sel, Nordbahnhofstr. la, 37213 Witzenhausen, Germany, E-mail: joerge@wiz.uni-kassel.de 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



304 R.G. Joergensen 

N mineralization is more strongly affected than C0 2 production (Nieder 
etal. 1993;Ahletal. 1998). 



■ Equipment 

• 100 mL low-density, wide-neck polyethylene (PE) bottles 

• Funnels 

• 50-mL Erlenmeyer flasks 

• Horizontal or overhead shaker 

• Folded filter papers (e.g., Whatman 42 or Schleicher & Schuell 595 1/2) 

• Room or incubator at 25° C 

• Steam distillation apparatus 

• Burette or autotitrator 

■ Reagents 

• 2 M KCl solution 

• 2%H 3 B0 3 (extra pure) 

• 10}xMHCl 

• MgO (extra pure) 

• Devarda alloy 

■ Sample Preparation 

Use field-moist, sieved (between < 2 and < 5 mm) soil at approx. 40-50% 
water holding capacity. 

■ Procedure 

1. Weigh 15 g field moist soil into nine PE bottles. 

2. Add 5 mL of water slowly. 

3. Incubate at 25 °C in the dark. 

4. Remove three replicates after 0, 14, and 28 days. 

5. Extract with 60 mL 2 M KCl (extractant-to-soil ratio of 4: 1 ) for 30 min by 
oscillating shaking at 200 rpm (or 45 min overhead shaking at 40 rpm). 



16 Determination of Aerobic N-Mineralization 305 

6. Filter through a folded filter paper. 

7. Pipette a 30 mL aliquot into the sample flask of the distillation appara- 
tus. 

8. Add approx. 200 mg MgO rapidly to volatize NH \ as NH 3 under alkaline 
conditions. 

9. Stop the first distillation when the distillate reaches the 30 mL mark 
on the receiver flask (a 50 mL Erlenmeyer flask containing 5 mL 2% 
H3BO3). 

10. Add approx. 200 mg Devarda alloy rapidly to reduce NO3 and NO2 to 
NHj, which is volatilized under the alkaline conditions of the distilla- 
tion flask. 

11. Stop the second distillation when the distillate reaches the 30 mL mark 
on the receiver flask (a 50-mL Erlenmeyer flask containing 5 mL 2% 
H3BO3). 

12. Titrate NH+ in each of the two distillates with 10 jim HC1 to pH 4.8. 



■ Calculation 

1. Calculation of extractable NH+ -N and NO3 -N 

xt/ / -n (S-B)xMxNx(VK + SW) 

N(ug/gsoil) = (16.1) 

^ && Ax DM 

S HC1 consumed by sample extract (pL) 

B HC1 consumed by blank extract (pL) 

M molarity of HC1 

N molecular mass of nitrogen (14) 

VK volume of K 2 S0 4 extractant (mL) 

SW total amount of water in the soil sample (mL) 

A sample aliquot (mL) 

DM total mass of dry soil sample (g) 

2. Calculation of net N mineralized 

N [|ig N/(g soil x day)] 

_ (NH+-N + NO" -N) t , +1 - (NH+-N + NO" -N) t , (16.2) 

n 



306 R.G. Joergensen 

td sampling day before the last sampling day (day or day 14) 
trf+i last sampling day (day 14 or day 28) 
n incubation period (days) 

■ Notes and Points to Watch 

• If the N mineralization rate of the first incubation period (0-14 days) 
does not differ significantly from that of the second incubation period 
(14-28 days), the average value of both periods should be used (Beck 
1983; Kandeler 1993a). If the N mineralization rate of the first incubation 
period is significantly lower than that of the second incubation period, 
e.g., due to N immobilization during the decomposition of plant residues, 
only the value of the second incubation period should be used. If the N 
mineralization rate of the first incubation period is significantly higher 
than that of the second incubation period, e.g., due to the increasing 
recalcitrance of decomposable soil organic matter, only the value of the 
first incubation period should be used. 

• The steam distillation method is especially suitable for colored extracts 
(Keeney and Nelson 1982; Forster 1995). 

• If a soil accumulates NO2 in the soil solution, a colorimetric method 
must be used to determine it (Keeney and Nelson 1982; Forster 1995). 

• Colorimetric methods are also available for the manual determination 
of extractable NO3 (e.g., Kandeler 1993a; Forster 1995), and for auto- 
mated segmented flow or flow injection, analyses are also available (e.g., 
Kutscha-Lissberg and Prillinger 1982). 

• It is possible to estimate the NO3 content in soil extracts by the decrease 
in UV absorbance after reduction of NO3 (Kandeler 1993b). 

• Colorimetric methods are also available for the manual determination of 
extractable NHJ (e.g., Keeney and Nelson 1982; Kandeler 1993; Forster 
1995), and for automated segmented flow or flow injection, analyses are 
also available. 

• Contamination of chemicals, especially KC1, but also of filter paper, 
funnel, extraction bottles, and glassware should be avoided and regularly 
checked. 

References 

Alef K (1995) Nitrogen mineralization in soils. In: Alef K, Nannipieri P (eds) Methods in 
Applied Soil Microbiology and Biochemistry. Academic Press, London, pp 234-245 



16 Determination of Aerobic N-Mineralization 307 

Ahl C, Joergensen RG, Kandeler E, Meyer B, Woehler V (1998) Microbial biomass and 
activity in silt and sand loams after long-term shallow tillage in central Germany. Soil 
Till Res 49:93-104 

Beck T (1983) Die N-Mineralisation von Boden im Brutversuch. Z Pflanzenernahr Bodenk 
146:243-252 

Forster JC (1995) Soil nitrogen. In: Alef K, Nannipieri P (eds) Methods in Applied Soil 
Microbiology and Biochemistry. Academic Press, London, pp 79-87 

Joergensen RG, Schmaedeke F, Windhorst K, Meyer B (1995) Biomass and activity of mi- 
croorganisms in a fuel oil contaminated soil. Soil Biol Biochem 27:1137-1143 

Kandeler E (1993a) Bestimmung der N-Mineralisation im aeroben Brutversuch. In Schin- 
ner F, Ohlinger R, Kandeler E, Margesin R (eds) Bodenbiologische Arbeitsmethoden, 
2nd ed. Springer, Berlin, pp 158-159 

Kandeler E (1993b) Bestimmung von Nitrat. In: Schinner F, Ohlinger R, Kandeler E, Mar- 
gesin R (eds) Bodenbiologische Arbeitsmethoden, 2nd ed. Springer, Berlin, pp 369-371 

Keeney DR, Nelson DW (1982) Nitrogen - inorganic forms. In: Page AL, Miller, RH, Kee- 
ner DR (eds) Methods of Soil Analysis, Part 2. Am Soc Agron, Soil Sci Soc Am, Madison, 
pp 643-698 

Kutscha-Lissberg P, Prillinger F (1982) Rapid determination of EUF-extractable nitrogen 
and boron. Plant Soil 64:63-66 

Nieder R, Scheithauer U, Richter J (1993) Dynamics of nitrogen after deeper tillage in arable 
loess soils of West Germany. Biol Fertil Soils 16:45-51 



17 



Determination of Enzyme Activities 
in Contaminated Soil 

Rosa Margesin 



17.1 

General Introduction 

Soil biological activities are sensitive to environmental stress; each change 
in environmental conditions may result in a shift in the species composition 
of the soil microflora and modification of their metabolic rate. Soil enzyme 
activities are attractive as indicators for monitoring various impacts on soil 
because of their central role in the soil environment. Soil enzymes are the 
catalysts of important metabolic processes including the decomposition of 
organic inputs and the detoxification of xenobiotics (Schinner et al. 1996; 
Dick 1997). 

Soil enzyme activities have been used as a biological indicator of pollu- 
tion with heavy metals, pesticides, and hydrocarbons (Schinner et al. 1993; 
Sparling 1997; van Beelen and Doelman 1997; Margesin et al. 2000a, 2000b). 
A number of studies have demonstrated that soil enzymes hold potential for 
assessing the impact of hydrocarbons and of fertilization on soil microor- 
ganisms and are a useful tool to monitor the early stages of remediation of 
contaminated soil (Margesin et al. 2000a, 2000b). The usefulness of various 
enzyme parameters depends on the composition and concentration of the 
hydrocarbons, as well as on other factors such as the age of contamination 
and physico-chemical soil characteristics. While some enzymes activities 
are appropriate to monitor the most active phase of biodegradation, oth- 
ers are also indicative of low hydrocarbon concentrations (Margesin et al. 
2000a). 

A broad spectrum of soil enzyme activities should be used to evaluate 
the effect of contamination on the different nutrient cycles. In this Chapter, 
a small selection of methods for the determination of enzyme activities 
in contaminated soil is described. Detailed descriptions of supplementary 
methods are given in Schinner et al. (1996). Of course, additional informa- 
tion on the soil biological status should be obtained from complementary 
methods, such as soil microbial counts (Chap. 13), soil biomass (Chap. 14), 
molecular biology (Chap. 10), and fatty acid profiles (Chap. 12). 



Rosa Margesin: Institute of Microbiology, Leopold Franzens University, Technikerstrasse 
25, 6020 Innsbruck, Austria, E-mail: rosa.margesin@uibk.ac.at 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



310 R. Margesin 

17.2 

Lipase-Esterase Activity 

■ Introduction 

Objectives. Soil lipase activity is a valuable tool to monitor the biodegrada- 
tion of petroleum hydrocarbons, such as diesel oil, in freshly contaminated 
soil (Margesin et al. 1999, 2002a, b). The residual hydrocarbon content 
correlates negatively with soil lipase activity in unfertilized as well as in 
fertilized soil (Margesin and Schinner 2001). Generally, a correlation be- 
tween soil lipase activity and other biological parameters can be found. 
Soil lipase activity increases with increasing initial oil loading rates, which 
demonstrates the induction of this enzyme activity by the contamina- 
tion. This induction is attributed to the appearance of products released 
from hydrocarbon biodegradation, which are the substrate for hydrolases 
including esterases-lipases. A strong relationship between the ability of mi- 
croorganisms to degrade diesel oil and their lipolytic activity was described 
by Mills et al. (1978) and Kato et al. (2001). No increase of lipase activity 
was observed during the biodegradation of polycyclic aromatic hydrocar- 
bons (PAHs) in soil (Margesin et al. 2000a). Assaying lipase activity is also 
useful to monitor the biodegradation of carboxyl esters such as lipids and 
biodegradable polyesters in soil (Sakai et al. 2002). The described method 
can also be applied to determine lipase activity in non-contaminated soil 
and could be useful for the screening of lipase-producing soil microorgan- 
isms. 

Principle. Using p-nitrophenyl butyrate (pNPB) as substrate, soil sam- 
ples are incubated at 30 °C and pH 7.25 for lOmin. After cooling on ice 
and centrifugation, the released p-nitrophenol (pNP) is determined spec- 
trophotometrically at 400 nm. To allow for the adsorption of pNP onto soil, 
a calibration curve is prepared in the presence of soil (Margesin et al. 2002). 

Theory. A significant proportion of lipids, such as pesticide emulsions, 
oils, and lipid conjugates, enter soil in the form of triacylglycerols, the 
primary storage fat in plant and animal tissue. The degradation of lipids is 
initiated by lipases (glycerol ester hydrolases) acting on the carboxylester 
bonds present in acylglycerols to liberate fatty acids and glycerol. Lipases 
are produced by a large variety of microorganisms, plants, and animals. 

The standard assays to determine the hydrolytic activity of lipase in 
soil are based on titration or fluorimetry. Methods that determine the 
fatty acids produced from tributyrin (Pokorna 1964; Hankin et al. 1982) 
or Tween 20 (Sakai et al. 2002) titrimetrically are easy to use, however, 
the disadvantages are the long incubation time (between 18 h and 3 days) 
and the possible adsorption of the released fatty acids onto soil colloids. 



1 7 Determination of Enzyme Activities in Contaminated Soil 311 

Fluorimetric methods (Pancholy and Lynd 1972; Cooper and Morgan 1981) 
are more sensitive and specific, but the substrate is relatively expensive. 
Colorimetric methods are quick and simple. Chromogenic substrates, such 
as p-nitrophenyl esters, are commonly used to assay microbial esterase and 
lipase activity (Shirai and Jackson 1982; Plou et al. 1998; Ishimoto et al. 
2001; Wei et al. 2003). In soil enzymology, p-nitrophenyl derivatives are 
widely used as substrates for measuring phosphatases, arylsulfatase, and 
6-glucosidase activity. The described method is based on the use of pNPB as 
substrate for the rapid, precise, and simple measurement of lipase activity 
in soil. 



■ Equipment 

• Centrifuge and centrifuge tubes (2,000 g, 2-4 °C) 

• Water bath (30 °C) 

• Spectrophotometer 

■ Reagents 

• Phosphate buffer: 100 mM NaH 2 P0 4 -NaOH, pH 7.25 (store at 4 °C) 

• Substrate: 100 mM p-nitrophenyl butyrate (pNPB) diluted in 2-propanol 
(store aliquots at -20 °C) 

• Calibration standards (p-nitrophenol, pNP) 

- Stock solution: 1 mg pNP/mL buffer (store at 4 °C) 

- Working standard solution: 100 yigpNP/mL buffer (prepare daily 
fresh) 

- Standards: 0, 25, 50, 75, 100, and 125 jig pNP/5 mL buffer (adjust vol- 
umes of 0-1.25 mL of the working standard solution to 5mL with 
buffer) 

■ Sample Preparation 

Use field-moist, sieved (< 5 mm) soil. 

■ Procedure 

1. Weigh 0.1 g of soil into centrifuge tubes, prepare 3-4 replicates (samples). 

2. Add 5 mL of buffer, and prewarm at 30 °C in a water bath for 10 min. 

3. Prepare a control (3-4 replicates) without soil. 

4. Add 50 p.L of substrate solution to each tube. 



312 R. Margesin 

5. Mix the contents and incubate the tubes in a water bath at 30 °C for 
exactly lOmin. 

6. Stop the reaction by cooling the tubes for 10 min on ice. 

7. Centrifuge the tubes at 2,000 g and 2-4 °C for 5 min. 

8. Pipette the supernatants in test tubes that are held on ice. 

9. Immediately afterward, measure the absorbance of the released pNP in 
samples and controls at 400 nm against the reagent blank. Dilute the 
solution with buffer when absorbance values are too high. 

To prepare a calibration curve, treat calibration standards like the soil 
samples: 

1. Weigh 0.1 g of soil into each of six centrifuge tubes. 

2. Add 5 mL of standard solution containing (= reagent blank), 25, 50, 75, 
100,andl25}igpNP. 

3. Mix and incubate at 30 °C for exactly 10 min. 

4. Proceed as described for the soil samples (6-8), and measure the ab- 
sorbance of the calibration standards at 400 nm against the reagent blank. 

■ Calculation 

1. Calculate the pNP concentration from the calibration curve. 

2. Subtract the control reading (hydrolysis in absence of soil) from the 
sample reading (hydrolysis in presence of soil) and express soil lipase 
activity as "|^g of released pNP per gram soil dry mass over 10 min" using 
the following formula: 

S-C 
|ig pNP/(g dry soil x 10 min) = 



wm x dm 



S pNP concentration of the soil sample (p.g) 
C pNP concentration of the control (pg) 
wm Soil wet mass (0.1 g) 
dm Soil dry mass (g) 

Notes and Points to Watch 

To measure the chemical pNP release from the substrate, it is necessary 
to prepare a control without soil. 



17 Determination of Enzyme Activities in Contaminated Soil 313 

• To allow for the adsorption of pNP onto soil, the calibration curve is 
prepared in the presence of soil. 

• It is important to use a neutral pH for this assay since the ester bond in 
pNPB is very labile and is fully hydrolyzed at alkaline pH. The degree 
of dissociation of pNP (colorless) into p-nitrophenoxide (yellow) is 0% 
below pH 5.0, 50% at pH 7.0 and 100% at pH 9.0. Enzyme assays based 
on p-nitrophenyl derivatives are usually carried out at a pH of 7.25 to 7.3 
(Ishimoto et al. 2001; Wei et al. 2003). 

• A reaction temperature of 30 °C is chosen to avoid a high rate of non- 
enzymatic substrate hydrolysis, which occurs at higher temperatures. 

• Drying and freeze-thawing of soil may affect soil lipase activity, however, 
this effect is soil-specific. 

• The presence of heavy metals in soil inhibits lipase activity. The metal 
sensitivity of soil lipases depends on the soil properties. 

• Lipase activity determined with the described assay has been found to 
correlate significantly with titrimetric and fluorimetric assays. 



17.3 

Fluorescein Diacetate Hydrolytic Activity 

■ Introduction 

Objectives. The rate of fluorescein diacetate (FDA) hydrolysis in soil has 
been considered a suitable index of overall enzyme activity (Schmirer and 
Rosswall 1982). It is a suitable indicator to indicate the onset of biodegra- 
dation of diesel oil and of monoaromatic compounds, such as BTEX (Mar- 
gesin et al. 2000a, 2003). The time course of FDA hydrolytic activity during 
the bioremediation of soil contaminated with diesel oil is comparable to 
that of lipase activity (Sect. 17.2). The biodegradation of PAHs (naphtha- 
lene, phenanthrene) in soil also results in an increase of FDA hydrolytic 
activity, which, however, is followed by a marked activity decrease (Mar- 
gesin et al. 2000a). A short-term, reversible inhibition of FDA hydrolysis 
has been noted in BTEX-contaminated soil (Margesin et al. 2003) and 
in jet-fuel-contaminated soil (Song and Bartha 1990), being substantially 
higher in unfertilized than in fertilized soil. The inhibition terminated after 
a significant part of the contamination had disappeared, and a stimulation 
of the activity was observed after most of the fuel had been mineralized. 
Generally, a correlation between FDA hydrolytic activity and other soil 
biological parameters can be found. 



314 R. Margesin 

Principle. Using FDA as substrate, soil samples are incubated at 25 °C and 
pH 7.6 for 2 h. The released fluorescein is extracted with acetone, and 
quantified photometrically at 490 nm. 

Theory. FDA is hydrolyzed by a number of different enzymes, such as pro- 
teases, lipases, and esterases. The ability to hydrolyze FDA is widespread 
among soil organisms and has been detected among heterotrophic bacteria, 
fungi, algae, and protozoa. The product of this enzyme conversion is flu- 
orescein, which can be visualized within cells by fluorescence microscopy. 
Fluorescein can also be quantified using fluorometry and spectrophotom- 
etry. Schmirer and Rosswall (1982) developed a simple, rapid, and sensitive 
spectrophometric method for the measurement of total microbial activity 
in soil straw litter. 

■ Equipment 

• Water bath (25 °C) 

• Spectrophotometer 

■ Reagents 

• Phosphate buffer: 60 mM NaH 2 P0 4 x H 2 0/Na 2 HP0 4 x 2H 2 0, pH 7.6 
(store at 4 °C) 

• Substrate solution: Fluorescein diacetate (FDA; 2 mg/mL) dissolved in 
acetone (store aliquots at -20 °C) 

• Acetone (technical grade) 

• Calibration standards (fluorescein) 

- Stock solution: 1 mg fluorescein/mL acetone (store at 4°C) 

- Working standard solution: 100 p.g fluorescein/mL buffer (prepare 
daily fresh) 

- Standards: 0, 20, 50, 70, 100, and 150p.g fluoresce in/ lOmL buffer 
(adjust volumes of 0-1.5 mL of the working standard solution to 
10 mL with buffer) 

■ Sample Preparation 

Use field-moist, sieved (< 5 mm) soil. 

■ Procedure 

1. Weigh 1 g of soil into 100 mL Erlenmeyer flasks, prepare 3 replicates 
(samples). 



17 Determination of Enzyme Activities in Contaminated Soil 315 

2. Add 10 mL of buffer. 

3. Prepare a control (3 replicates) without soil. 

4. Add 100 pi of substrate solution to each flask. 

5. Mix the contents and incubate the stoppered flasks at 25 °C for 2 h. 

6. Stop the reaction by adding 10 mL of acetone. 

7. Filter the contents of the flasks. 

8. Immediately afterwards, measure the absorbance of the released fluo- 
rescein in samples and controls at 490 nm against the reagent blank. 

To prepare a calibration curve, treat calibration standards like the soil 
samples: 

1. Weigh 1 g of soil into each of six flasks. 

2. Add 10 mL of standard solution containing (= reagent blank), 20, 50, 
70, 100, 120, and 150 pg fluorescein. 

3. Proceed as described for the soil samples (5-7), and measure the ab- 
sorbance of the calibration standards at 490 nm against the reagent blank. 

■ Calculation 

3. Calculate the fluorescein concentration from the calibration curve. 

4. Subtract the control reading (hydrolysis in absence of soil) from the 
sample reading (hydrolysis in presence of soil) and express activity as 
"|^g of released fluorescein per gram soil dry mass over 2 h" using the 
following formula: 

S-C 
|ig Fluorescein/(g dry soil x 2 h) = 



wm x dm 



S pNP concentration of the soil sample (pg) 
C pNP concentration of the control (pg) 
wm Soil wet mass (1 g) 
dm Soil dry mass (g) 

Notes and Points to Watch 

To measure the chemical fluorescein release from the substrate, it is 
necessary to prepare a control without soil. 



316 R. Margesin 

• To allow for the adsorption of fluorescein onto soil, the calibration curve 
is prepared in the presence of soil. This is especially important in case of 
soil containing high amounts of organic matter or clay. 

• Depending on the soil to be tested it might be necessary to optimize soil 
mass, substrate concentration, and incubation time. 

• It is important to use a neutral pH for this enzyme assay since chemical 
hydrolysis of FDA occurs under acidic and alkaline pH conditions. 

• Chemical hydrolysis of FDA occurs also at higher temperatures, therefore 
a reaction temperature of 25 °C is used for this enzyme assay. Store 
aliquots of the substrate solution at -20 °C to avoid chemical substrate 
hydrolysis, which occurs when the solution is stored for longer periods 
at 4 °C or room temperature. 

• Acetone is added not only to stop the enzyme reaction, but also to solu- 
bilize cell membranes in order to facilitate the extraction of fluorescein 
from microbial cells. 

• High amounts of heavy metals in soil may interfere with the method. 



17.4 

Dehydrogenase Activity 

■ Introduction 

Objectives. Soil dehydrogenase activity is a useful method to monitor the 
bioremediation of soil contaminated with petroleum hydrocarbons, such 
as diesel oil. The method has been applied to soil containing fresh (Mar- 
gesin and Schinner 1997; Margesin et al. 2000a) and aged contamination 
(Margesin and Schinner 1999, 2001). A statistically significant positive cor- 
relation between this activity and the residual hydrocarbon content has 
been repeatedly observed. The substantial increase in soil dehydrogenase 
activity after hydrocarbon contamination reflects the adaptation and ex- 
ponential growth of hydrocarbon degraders due to the availability of new 
carbon sources introduced by the contamination. Soil dehydrogenase activ- 
ity declines with decreasing hydrocarbon content due to the loss of available 
compounds as a consequence of biodegradation (Margesin et al. 2000a, b). 
Measuring dehydrogenase activity is also a useful tool to monitor envi- 
ronmental contamination by anionic surfactants (Margesin and Schinner 
1998), since this activity is inhibited in presence of high concentrations 
of anionic surfactants, such as sodium dodecyl sulfate. However, dehydro- 
genase activity is not a suitable parameter to monitor biodegradation of 
PAHs in soil (Margesin et al. 2000a). 



17 Determination of Enzyme Activities in Contaminated Soil 317 

Principle. Soil samples are mixed with [2(p-iodophenyl)-3-(p-nitrophe- 
nyl)-5-phenyl tetrazolium chloride] solution (INT), and incubated for 2h 
at 40 °C. The reduced iodonitro tetrazolium formazan (INTF) is extracted 
with dimethylformamide and ethanol, and quantified photometrically at 
464 nm (von Mersi and Schinner 1991; Schinner et al. 1996). 

Theory. Dehydrogenases belong to oxidoreductases and catalyze the re- 
moval from a substrate of two hydrogens that are taken up by a hydrogen 
acceptor or coenzyme. Nicotinamide adenine dinucleotide (NAD) is the 
coenzyme used by the majority of dehydrogenases, others use nicotinamide 
adenine dinucleotide phosphate (NADP) or flavin adenine dinucleotide 
(FAD). Since dehydrogenases are important components of the enzyme 
system of all microorganisms, soil dehydrogenase activity reflects a broad 
range of microbial oxidative activities, and can be taken as a measure for the 
intensity of microbial metabolism in soil (Schinner et al. 1996). Because of 
increased sensitivity and reproducibility, the substrate INT has been used 
by a number of authors (Trevors 1984; Griffiths 1989; von Mersi and Schin- 
ner 1 99 1 ) to determine soil dehydrogenase activity. An alternative substrate 
is 2,3,5-triphenyltetrazolium chloride (for details, see Schinner et al. 1996). 

■ Equipment 

• Water bath or incubator (40 °C) 

• Spectrophotometer 

■ Reagents 

• Buffer: 1 M Tris-HCl, pH 7.0 (store at 4 °C). 

• Substrate: dissolve 500 mg of 2(p-iodophenyl)-3-(p-nitrophenyl)-5-phe- 
nyl tetrazolium chloride (INT; Serva, Heidelberg) in 2 mL of N,N-dime- 
thylformamide by shaking vigorously and using an ultrasonic water 
bath. Make up to volume with distilled water in a 100 mL volumetric 
flask (prepare daily fresh, and store it in the dark until use). 

• Extraction solution: mix 1 part of N,N-dimethylformamide with 1 part 
of 96% ethanol. 

• Calibration standards (iodonitrotetrazolium formazan (INTF)). 

- Stock solution: 100 yig INTF/mL extraction solution (store at 4 °C) 

- Standards: 0, 100, 200, 300, and 500 jig INTF/13.5 mL extraction solu- 
tion (adjust volumes of 0-5 mL of the stock solution to 13.5 mL with 
extraction solution) 



318 R. Margesin 

■ Sample Preparation 

Use field-moist, sieved (< 5 mm) soil. Additionally, prepare autoclaved soil. 

■ Procedure 

1. Weigh 1 g of soil into 100 mL Erlenmeyer flasks, prepare 3 replicates 
(samples). 

2. Weigh 1 g of autoclaved soil into 100 mL Erlenmeyer flasks, prepare 2 
replicates (controls). 

3. Add 1.5 mL of buffer and 2 mL of substrate solution to both samples and 
controls. 

4. Mix the contents and incubate the stoppered flasks at 40 °C for 2 h. 

5. Add lOmL of extraction solution to each flask. For extraction of the 
released INTF keep the flasks for 1 h at room temperature in the dark, 
and shake vigorously every 20 min. 

6. Filter the contents of the flasks. 

7. Immediately afterwards, measure the INTF concentration in samples, 
controls and calibration standards at 464 nm against the reagent blank. 
Dilute solutions with extraction solution when absorbance values are 
too high. 

■ Calculation 

1. Calculate the INTF concentration from the calibration curve. 

2. Express soil dehydrogenase activity as "|ig of released INTF per gram 
soil dry mass over 2 h" using the following formula: 

S-C 
|Lig INTF/(g dry soil x 2 h) = 



wm x dm 



S INTF concentration of the soil sample (p.g) 
C INTF concentration of the control (p.g) 
wm Soil wet mass (1 g) 
dm Soil dry mass (g) 

Notes and Points to Watch 

INT is very sensitive to light. Both incubation and filtration have to be 
carried out in the dark. 



17 Determination of Enzyme Activities in Contaminated Soil 319 

• The control contains autoclaved soil to estimate the chemical substrate 
reduction due to reactive soil components. 

• Depending on the soil to be tested, it might be necessary to optimize soil 
mass, substrate concentration, and incubation time. 

• High amounts of heavy metals (e.g., copper) in soil can interfere with 
the method. 

• Dehydrogenase activity is significantly reduced in acidic soils. 

References 

Cooper AB, Morgan HW (1981) Improved fluorimetric method to assay for soil lipase 

activity. Soil Biol Biochem 13:307-311 
Dick RP (1997) Soil enzyme activities as integrative indicators of soil health. In: Pankhurst 

CE, Double BM, Gupta VV (eds) Biological indicators of soil health. CAB Int, Oxon, 

pp 121-157 
Griffiths BS (1989) Improved extraction of iodonitrotetrazolium-formazan from soil with 

dimethylformamide. Soil Biol Biochem 21:179-180 
Hankin L, Hill DE, Stephens GR (1982) Effect of mulches on bacterial populations and 

enzyme activity in soil and vegetable yields. Plant & Soil 64:193-201 
Ishimoto R, Sugimoto M, Kawai F (2001) Screening and characterization of trehalose-oleate 

hydrolyzing lipase. FEMS Microbiol Lett 195: 231-235 
Kato T, Haruki M, Imanaka T, Morikawa M, Kanaya S (2001) Isolation and characteriza- 
tion of psychrotrophic bacteria from oil-reservoir water and oil sands. Appl Microbiol 

Biotechnol 55:794-800 
Margesin R, Feller G, Hammerle M, Stegner U, Schinner F (2002) A colorimetric method 

for the determination of lipase activity in soil. Biotechnol Lett 24:27-33 
Margesin R, Schinner F (1997) Bioremediation of diesel-oil-contaminated alpine soils at low 

temperatures. Appl Microbiol Biotechnol 47:462-468 
Margesin R, Schinner F (1998) Biodegradation of the anionic surfactant sodium dodecyl 

sulfate at low temperatures. Int Biodet Biodegradation 41:139-143 
Margesin R, Schinner F (1999) A feasibility study for the in situ remediation of a former 

tank farm. World J Microbiol Biotechnol 15:615-622 
Margesin R, Schinner F (2001) Bioremediation (natural attenuation and biostimulation) of 

diesel-oil-contaminated soil in an alpine glacier skiing area. Appl Environ Microbiol 

67:3127-3133 
Margesin R, Walder G, Schinner F (2000a) The impact of hydrocarbon remediation (diesel oil 

and polycyclic aromatic hydrocarbons) on enzyme activities and microbial properties 

of soil. Acta Biotechnol 20:313-333 
Margesin R, Walder G, Schinner F (2003) Bioremediation assessment of a BTEX- 

contaminated soil. Acta Biotechnol 23:29-36 
Margesin R, Zimmerbauer A, Schinner F (1999) Soil lipase activity - a useful indicator of 

oil biodegradation. Biotechnol Tech 13:859-863 
Margesin R, Zimmerbauer A, Schinner F (2000b) Monitoring of bioremediation by soil 

biological activities. Chemosphere 40:339-346 
Mills AL, Breuil C, Colwell RR (1978) Enumeration of petroleum-degrading marine and 

estuarine microorganisms by the most probable number method. Can J Microbiol 

24:552-557 



320 R. Margesin 

Pancholy SK, Lynd JQ (1972) Quantitative fluorescence analysis of soil lipase activity. Soil 

Biol Biochem 4: 257-259 
Plou FJ, Ferrer M, Nuero OM, Calvo MV, Alcalde M, Reyes F, Ballesteros A (1998) Analysis of 

Tween 80 as an esterase/lipase substrate for lipolytic activity. Biotechnol Tech 12: 183- 186 
Pokorna V (1964) Method of determining the lipolytic activity of upland and lowland peats 

and muds. Soviet Soil Sci 1:85-87 
Sakai Y, Hayatsu M, Hayano K (2002) Use of Tween 20 as a substrate for assay of lipase 

activity in soils. Soil Sci Plant Nutr 48:729-734 
Schinner F, Bayer H, Mitterer M (1993) The influence of herbicides on microbial activity in 

soil materials. Austrian J Agric Res 34:22-30 
Schinner F, Ohlinger R, Kandeler E, Margesin R (eds; 1996) Methods in Soil Biology. Springer, 

Berlin Heidelberg New York 
Schniirer J, Rosswall T (1982) Fluorescein diacetate hydrolysis as a measure of total microbial 

activity in soil and litter. Appl Environ Microbiol 43:1256-1261 
Shirai K, Jackson RL (1982) Lipoprotein lipase-catalyzed hydrolysis of p-nitrophenyl bu- 

tyrate. J Biol Chem 257:1253-1258 
Song HG, Bartha R (1990) Effects of jet fuel on the microbial community of soil. Appl 

Environ Microbiol 56:646-651 
Sparling GP (1997) Soil microbial biomass, activity and nutrient cycling as indicators of 

soil health. In: Pankhurst CE, Double BM, Gupta VV (eds) Biological indicators of soil 

health. CAB Int, Oxon, pp 97-119 
Trevors JT (1984) Dehydrogenase activity in soil: A comparison between the INT and TTC 

assay. Soil Biol Biochem 16:673-674 
van Beelen P, Doelman P (1997) Significance and application of microbial toxicity tests in 

assessing ecotoxicological risks of contaminants in soil and sediment. Chemosphere 

34:455-499 
Von Mersi, Schinner F (1991) An improved and accurate method for determining the 

dehydrogenase activity of soils with iodonitrotetrazolium chloride. Biol Fertil Soils 

11:216-220 
Wei YL, Kurihara T, Suzuki T, Esaki N (2003) A novel esterase from a psychrotrophic 

bacterium, Acinetobacter sp. strain no. 6, that belongs to the amidase signature family. 

J Mol Cat B-Enzym 23:357-365 



^ O Assessment of Ecotoxicity 
' ^ of Contaminated Soil Usin 



Using Bioassays 

Adolf Eisentraeger, Kerstin Hund-Rinke, Joerg Roembke 



18.1 

General Introduction: Strategy 

Bioassays provide important information for the assessment of pollutant 
effects of chemicals or environmental samples. In contrast to chemical 
analyses, they also detect effects of multiple contaminants and metabo- 
lites. Standardized bioassays can be used for the path-related, toxicological 
characterization of soils and soil materials, taking into account possible 
transfer of pollutants to the groundwater and potential effects on soil mi- 
croorganisms, earthworms, and plants. A large number of bioassays have 
been applied for the characterization of contaminated soil or soil materials 
(Spurgeon et al. 2002). Most of them have been developed for the testing 
of chemicals and then have been adapted for testing of contaminated soil 
samples. In the first case, uncontaminated soil is spiked with chemicals in 
defined concentrations and dose-response relationships are obtained and 
evaluated further. While this is a straightforward and often standardized 
approach, testing and assessing contaminated soils is more difficult, since 
many soil samples are contaminated with different kinds of known and un- 
known chemicals that can not be quantified comprehensively. In addition, 
the soils have different properties (e.g., pH, texture), which themselves can 
affect organisms. Whereas the uncontaminated soil sample can be used 
as reference sample for the testing of chemicals, it is very difficult to se- 
lect uncontaminated reference samples for contaminated soils. Therefore, 
bioassays can not be transferred easily to the testing of contaminated soils 
and the evaluation of test results is completely different. 

The results of bioassays using soil are affected by mobilization, bioavail- 
ability, and pathways of transfer of contaminants. The latter ones can be 
varied by the kind of organisms and the test design chosen. In addition, 
by using a battery of different test systems the effects of contaminants can 
be assessed as a whole (i.e., whether they are known or not), thus covering 

Adolf Eisentraeger: Institute of Hygiene and Environmental Medicine, Aachen University of 

Technology, Pauwelsstr. 30, 52074 Aachen, Germany, E-mail: adolf.eisentraeger@post.rwth- 

aachen.de 

Kerstin Hund-Rinke: Fraunhofer Institute for Molecular Biology and Applied Ecology, P.O. 

Box 1260, 57377 Schmallenberg, Germany 

Joerg Roembke: ECT Oekotoxikologie GmbH, Boettgerstr. 2-14, 65439 Floersheim, Germany 



Soil Biology, Volume 5 

Manual for Soil Analysis 

R. Margesin, F. Schinner (Eds.) 

© Springer- Verlag Berlin Heidelberg 2005 



322 A. Eisentraeger et al. 

potential synergistic or antagonistic effects. Therefore, bioassays are useful 
tools for complementing chemical analysis. 

Several joint research projects were carried out in the recent years in 
order to optimize bioassays with respect to sample treatment, test perfor- 
mance, and evaluation of data (Hund-Rinke et al. 2002a, 2002b). After per- 
forming a round-robin test and a laboratory intercomparison test, a testing 
strategy was proposed by researchers involved in these projects (Dechema 
2001; Eisentraeger et al. 2004). The strategy allows a cost efficient assess- 
ment of soil samples in addition to chemical analysis in a stepwise approach 
using a maximum of nine standardized bioassays. Detailed advice both on 
sample treatment and on the interpretation of test data is given. Depending 
on the results, recommendations are given whether remediated soils can 
be incorporated at unsealed or sealed sites and whether they can be used as 
upper or lower soils. The proposed strategy can be expected to contribute 
to the discussion on the standardization of soil testing and to promote the 
incorporation of biological test methods into soil protection legislation. 

As mentioned above, nine different bioassays are applied in a stepwise 
approach. The four pathways from soil to (ground)water, to soil microflora, 
to soil fauna and higher plants are assessed using several test systems. 

The water-extractable ecotoxicological potential of soils and soil mate- 
rials is examined to assess whether undesirable effects in the groundwater 
or surface water might occur (i.e., after toxic substances have been set free) 
by using two aquatic ecotoxicity tests with bacteria and algae (Table 18.1). 
Since only the potential is of significance in this context, the selected test 
systems are not considered to be ecologically relevant. 

Genotoxic substances in contaminated soils may be hazardous both for 
soil organisms and human beings. The latter may be exposed via the path 
soil - groundwater - since it is one of the major sources for drinking water. 
Therefore, the water extractable genotoxic potential is assessed by testing 
water extracts using a test of genotoxicity known as the umu test (because 
of its dependence on umuC gene induction) according to ISO 13829 (2000; 
Table 18.1). The Salmonella! 'microsome test (Ames test) according to DIN 
38415 T4 (1999) should be carried out additionally, but only if the umu test 
is negative and there are strong hints from chemical analysis or site history 
that mutagenic compounds are present (Table 18.1). It should be noted that 
the approach presented here is a screening method to identify substances 
that can cause gene mutations; it cannot be used to identify clastogenic 
substances. 

Several bioassays are employed to ascertain different aspects of the habi- 
tat function of soils (Table 18.2): Microorganisms are chosen for these that 
differ in trophic levels, exposition, and habitat (e.g., bulk soil, air- filled soil 
pores; water film of soil pores). Effects on the soil microflora are quantified 
via respiration and ammonium oxidation activity. The combined earth- 



18 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 



323 



Table 18.1. Biological test systems for the ecotoxicological assessment of the water- 
extractable ecotoxic and genotoxic potential of contaminated soil or soil materials. (Eisen- 
traeger et al. 2004) 



Test system 



Standard 



Comments/modifications 



Water quality - determination 
of the inhibitory effect of water 
samples on the light emission 
of Vibrio fischeri luminescent 
bacteria test - parts 1-3 

Water quality - fresh water al- 
gal growth inhibition test with 
Scenedesmus subspicatus and 
Selenastrum capricornutum 

Water quality - determination 
of the genotoxicity of water 
and waste water using the umu 
test 



ISO 11348 (1998) 



ISO 8692 (1989) 



ISO 13829 (2000) 



Alternatively testing on 

microplates possible 

(Eisentraeger et al. 2003; 

Rilaetal.2003) 



Water quality - determina- 
tion of the genotoxicity of wa- 
ter and waste water using the 
Salmonella/ microsome test 
(Ames test) 



DIN 38415 T4 (1999) 



worm mortality/reproduction test and the Collembola reproduction test 
are used to assess effects on soil fauna. Further, the emergence and growth 
of Brassica rapa and Avena sativa are used to assess a soil's capacity to 
function as a habitat for higher plants. 



18.2 

Sample Preparation 

■ Introduction 

Objectives and Principles. Suitable sample preparation is a prerequisite for 
obtaining reliable results (ISO 15799 2003). The preparation of soil sam- 
ples includes transport, sieving, determination of maximum water-holding 
capacity, and water content, as well as adjustment of water content and stor- 
age. 

Theory. Tests should be performed as soon as possible after sampling. The 
period of storage should be minimized, at least for soils containing degrad- 



324 



A. Eisentraeger et al. 



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1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 325 

able contaminants. Soil sampling may cause an alteration of the soil con- 
ditions, especially of the redox potential. This may result in a degradation 
of the contaminants and lead to an inaccurate evaluation of toxic potential. 
Due to altered environmental conditions, for example, it was found at a mu- 
nition site that the TNT concentration after excavation quickly decreased 
by a factor of about 10. 

Microbial activity may diminish in storage, leading to erroneous results 
with respect to microbial respiration and nitrification rates (see below). 
The extent and time period of decrease varies for different soils. If no 
degradation is expected, the maximum storage period for uncontaminated 
soil samples should not exceed 3 months at 4 °C in the dark. If soil samples 
have to be stored for a longer period, -20 °C may be best. Certain aspects 
of these procedures, however, remain controversial, especially regarding 
potential ammonium oxidation activity (Sect. 18.5.2). 

Samples should be stored in a way that changes in the soil water content 
are minimized. The vessels used should not influence the composition of the 
samples. For soil samples contaminated with organic pollutants, stainless 
steel, aluminum, or glass vessels should be used. Container materials of 
lower quality may be used for large amounts of soil, in which case it must 
be assured that the ratio of soil mass to vessel mass is appropriate. The 
decrease of contaminant concentrations (e.g., by sorption to the wall of the 
vessel) also must be negligible. Suitable containers for samples containing 
inorganic contaminants (heavy metals) usually made of plastic and or other 
materials free of heavy metals. 

A draft version of a sequential approach to estimate the storage capacity 
of soil samples contaminated with volatile organic compounds (VOC) was 
set up by Rila and Eisentraeger (2003). This approach is based on the quan- 
tification of the VOCs, on the one hand, and of the microbial respiratory 
activity, on the other, under the assumption that toxic characteristics of 
samples with a high microbial activity and a high VOC concentration are 
altered during storage. 

The procedure of sample preparation is summarized in Fig. 18.1. Ac- 
cording to ISO/DIS 21268 (2004), for the tests with soil eluate, and for those 
using terrestrial test organisms other than microorganisms, the soil sam- 
ples are sieved to < 4 mm. For microbial tests the soil fraction < 2 mm 
is needed. The microbial tests are performed using the indigenous soil 
microflora, whereas the other tests are performed with introduced organ- 
isms. These latter tests usually require huge amounts of soil. Especially for 
highly silty and loamy soils, sieving of large soil volumes to smaller soil 
fractions may be difficult with an acceptable expenditure of work, as the 
holes of the sieves may plug up within several minutes. This makes frequent 
cleaning necessary. Therefore, depending on the organisms introduced, it 
was decided to apply different procedures. 



326 



A. Eisentraeger et al. 



SOIL SAMPLING, transportation, storage, 
documentation of soil treatment 



I 



MAIN SIEVING 

(< 4 mm nominal screen size; soil sample 

of at least 2 kg), quantification of water 

content and water holding capacity 



i 



WATER EXTRACTION 

using a glass flask (500 ml) with defined 

amount of soil (175 g ± 5 g), addition of 

water (soil / water 1+2) 

AGITATION for 24 h: 

end-over-end tumbler: 5-10 rpm or 

roller table rotating at about 10 rpm 



I 



SETTLING 

of suspended soils (15 ± 5 min) 



I 



CENTRIFUGATION 

at 2,000 - 3,000 g for 20 min 



I 



FILTRATION a > 

(0.45 um glass fibre membrane filter 

without organic glue or regenerated 

cellulose) 



I 



BIOASSAYS: 

water extractable ecotoxicity and 
genotoxicity 



I 



Adjustment of the water content to 
50 ± 10 % WHCmax 

INVESTIGATION OF PATHWAYS 
SOIL - SOIL FAUNA and PLANTS 



Sieving < 2 mm; adjustment of the water 
content to 50 ± 10 % WHCmax 

INVESTIGATION OF PATHWAY 
SOIL - SOIL MICRO-ORGANISMS 



Centrifugation at 10,000 - 20,000 g b > 



if no genotoxic potential: 
solid phase extraction of water extract 



I 



BIOASSAYS: 

water extractable genotoxicity 



a ) As the standard can be used for 
the determination of organic pol- 
lutants in the eluate and for its 
ecotoxicological characterization, 
it must be kept in mind that the 
use of 0.45 yam filters may reduce 
both the concentration of organic 
pollutants and the toxicity because 
pollutants adsorbed to soil parti- 
cles will be blocked by the filter. 

b ) If the concentration of bound 
pollutants or the global toxicity 
of the eluate are of interest other 
techniques such as settling or cen- 
trifugation are recommended. 



Fig. 18.1. Procedure proposed for the preparation of soil samples for ecotoxicological testing 
(modified according to Pfeifer et al. 2000; Dechema 2001; Rila and Eisentraeger 2003; 
Eisentraeger et al. 2004, ISO/DIS 21268-1 2004) 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 327 

Sample Preparation for Investigation of the Pathways: Soil to-Soil Organisms, and Soil 
to Plants. It has to be considered that biological determinations require 
optimal water content. The water content also has an influence on the 
oxygen supply; water saturation, for example, results in a limited oxy- 
gen supply. As the demands of organisms regarding humidity and oxygen 
supply differ, the optimal water content depends on the planned investi- 
gations. For microbiological assessments a water content of 50 ± 10% of 
the maximum water holding capacity (WHC max ) is recommended. Soils 
with this water content provide the microorganisms living in the soil pore 
water with sufficient water as well as oxygen. Similar conditions are rec- 
ommended for collembola that live in air-filled soil pores. For tests with 
earthworms the appropriate water content is higher, since their surface is 
rather sensitive to desiccation. Due to their large size, the thin water film in 
soil pores, sufficient for microorganisms, is insufficient for them. Further- 
more, the earthworm species used in ecotoxicological tests is a compost 
worm, adapted to higher humidity. 

The soil has to be dried if the water content is too high for the planned 
investigations or for sieving. During the drying process local complete 
drying should be avoided. This is essential for microbial investigations, 
since complete drying causes a reduction of the microbial population. 
Moreover, the aggregates formed in silty and loamy soils will be difficult to 
destroy. Localized drying can be prevented by turning the soil periodically. 
In addition, structural changes of the soil due to drying can cause problems 
in plant tests since roots cannot permeate hard soil aggregates. In such 
cases, the soil samples must be re-wetted carefully by hand in order to 
reach a moisture suitable for plants, i.e., automatic moistening via wicks 
(Sect. 18.5.5) will not be sufficient. 

Water Extraction for Ecotoxicological and Genotoxicological Testing. In order to 
make a pragmatic estimation of the fraction of contaminants that might 
migrate to the groundwater, soil samples are extracted with water in a simple 
batch assay. By choosing a dry soil-to-water ratio of 1:2 it is guaranteed, 
on the one hand, that enough water sample is available for biotesting and, 
on the other, that concentrations of the water-extractable contaminants 
remain high. Meanwhile, this approach is accepted, has been successfully 
validated in a ring test, and its standardization is in progress (ISO/DIS 
21268-1 2004). 

Preparation of Solid-Phase Extracts from the Water Extracts for Genotoxicological 
Testing. As stated earlier, groundwater contaminated with genotoxic sub- 
stances can be hazardous. The water-extractable genotoxic potential is 
assessed in order to roughly estimate whether genotoxic compounds might 
be mobilized by water. In the first step of the procedure, the same water 
extract is tested as is used for the assessment of the water-extractable eco- 



328 A. Eisentraeger et al. 

toxicological potential. If there is a genotoxic effect in the umu test, with 
or without metabolic activation, a high risk of transfer of genotoxic sub- 
stances from soil to groundwater exists. If there is no genotoxic effect, the 
water extract should be concentrated by a (low) factor of 15 using Serdolit 
PAD-1 resin (Boehringer Ingelheim Pharma GmbH & Co. KG, Ingelheim, 
Germany) as described below. 

■ Equipment and Reagents 

• 2 mm sieve (in exceptional cases 4 mm, see Sect. 18.1) 

• Cylinders of metal, glass, or plastic (diameter 5-8 cm), sealed at one end 
with a finely meshed fabric for determining water-holding capacity 

• Sand bath fitted out at the bottom with a discharge valve, filled with fine 
sand (grain size 0.1-0.7 mm); to about 10 cm, then saturated with water 
before starting the test by closing the valve while letting in the water and 
opening it afterward (so that the surplus water can run off), and the sand 
then covered with a moist fabric 

• Analytical balance 

• Heating apparatus for determining water content or drying cabinet and 
exsiccator 

• Shakers for water extraction 

• Centrifuge 

• Glass microfiber filters 

• Pentane, acetone, dimethylsulfoxide (DMSO), methanol, dichlorome- 
thane, cone. HC1, cone. NaOH (of analytical grade) 

• Resin (e.g., Serdolit PAD-1 resin; Boehringer Ingelheim, No. 42442) 

■ Sample Preparation 

The preparation of the soil consists of the following steps: 

• Sieving 

• Water extraction for aquatic test systems 

• Solid phase extraction of the water extract for aquatic genotoxicity test 
systems 

• Determination of the WHC max (Chap. 2) 

• Determination of the water content of the sieved soil (Chap. 2) 

• Adjustment of the water content to a specific percentage of WHC 
(Chap. 2) 



max 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 329 

■ Procedure 

In the guideline ISO 10381-6 (1993) collection, handling, and storage of 
soil for the assessment of aerobic microbial processes in the laboratory is 
described. For testing contaminated soils it has to be considered that some 
contaminants may interact with vessel material (see Sect. 18.1). Moreover, 
alteration of the redox potential during storage should be minimized for 
anaerobic soils for which only investigation by aquatic ecotoxicological and 
genotoxicological tests is relevant. 

Sieving (According to ISO 10381-6 1993) 

If the soil is too wet for sieving, it should be spread out, where possible in 
a gentle air stream, to facilitate uniform drying. The soil should be finger 
crumbled and turned over frequently to avoid excessive surface drying. 
Normally this procedure should be performed at ambient temperature. 
The soil should not be dried more than necessary to facilitate sieving. 

Water Extraction (According to ISO/DIS 21268-2 2004) 

The soil samples are extracted by a ratio of 1 part soil dry mass to 2 parts of 
water with a minimum amount of 100 g soil dry mass. The water content in 
the soil has to be considered. The samples are shaken intensively to simulate 
worst-case conditions for 24 h and then centrifuged. The supernatant is 
filtered with a glass microfiber filter and stored at 4°C in Duran (Schott 
AG, Mainz) glass bottles in the dark. The pH of the elutriates is adjusted 
to 7 ± 1 with cone. HC1 or NaOH. Ecotoxicological and genotoxicological 
testing should be performed within 8 days. 

Preparation of Solid-Phase Extracts from the Water Extracts 
for Genotoxicological Testing 

The solid-phase extraction of the water extract is performed with Serdo- 
lit PAD-1 resin, an ethylstyrene-DVB-copolymer with a particle size of 
0.3-1.0 mm and a pore diameter of ca. 25 nm with a specific surface of 
ca. 250m 2 /g. The PAD-1 beads are pretreated by rinsing for 2h in warm 
10% (v/v) HC1, Millipore water, 10% (v/v) NaOH, and Millipore water 
successively followed by 8h Soxhlet extraction with pentane/acetone in 
a ratio of 1:2. The beads are dried at a temperature of 1 10 °C. Shortly before 
solid phase extraction 10 g PAD-1 beads are preconditioned by shaking 
them with 25 mL methanol. 

The water extract should be concentrated by a factor of 15 by mixing 
375 mL with 10 g Serdolit PAD-1 beads. This suspension is placed on an 
overhead shaker for 2.5 h. The beads are removed from the water extract 
and dried under nitrogen atmosphere in a Baker-spe-10 system (J.T. Baker, 



330 A. Eisentraeger et al. 

Phillipsburg, New Jersey, USA). The dried beads are then extracted with 
a mixture of 9 parts dichloromethane and 1 part methanol. One mL of 
DMSO is added to the solvent, which is then evaporated under nitrogen 
atmosphere to a final volume of 1 mL. The concentrated sample is stored 
for less than 8 days at 4°C. The sample is adjusted with distilled water to 
a volume of 25 mL for the genotoxicity tests. The final DMSO concentration 
is 4%. Therefore, the concentration factor for the water soil extract is 15. 

■ Notes and Points to Watch 

• As already mentioned in Sect. 18.1, localized drying of the soil has to be 
avoided. 

• The soil should be processed as soon as possible after sampling. Any 
delays due to transportation should be minimized. 

• Microbial tests: if storage is unavoidable, this should not exceed 3 months, 
unless evidence of continued microbial activity is provided. Even at low 
temperatures the active soil microflora decreases with increasing storage 
time; the rate of decrease depends on the composition of the soil and the 
microflora. 

• Soil fauna tests and tests using higher plants: there are no specific recom- 
mendations for soil storage with respect to soil fauna and higher plants 
in ISO standards. Therefore it is recommended to store the soil sam- 
ples under the same conditions as for testing of microbes and microbial 
processes. 

• Aquatic tests: for testing the leaching potential, water extracts for aquatic 
tests should be prepared immediately after sieving. If the tests cannot be 
performed within 8 days (storage of the extracts at 4 ± 2 °C in the dark), 
extracts should be stored at -20 °C. 

• An ISO guidance paper on the long and short term storage of soil samples 
is in process. 

18.3 

Water-Extractable Ecotoxicity 

18.3.1 

Vibrio fischeri Luminescence-Inhibition Assay 

■ Introduction 

Objectives. This test is an acute toxicity test with the marine lumines- 
cent bacterium Vibrio fischeri NRRL B-11177 (formerly known as Photo- 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 331 

bacterium phosphoreum). It is standardized for the determination of the 
inhibitory effect of water samples in the ISO guideline 11348 parts 1-3 
(1998). In the strategy presented here, it is used to determine whether toxic 
substances are present in the aqueous soil extracts. 

Principle. The test system measures the light output of the luminescent 
bacteria after they have been challenged by a sample and compares it to 
the light output of a blank control sample. The difference in light output 
(between the sample and the control) is attributed to the effect of the sample 
on the organisms. The test is based on the fact that the light output of the 
bacteria is reduced when it is introduced to toxic chemicals. 

Theory. V.fischeri emits a part of its metabolic energy as blue-green light 
(490 nm). Biochemically luminescence is a byway of the respiratory chain. 
Reduction equivalents are separated and transmitted to a special acceptor 
(flavin mononucleotide, FMN; Engebrecht et al. 1983). During the oxidation 
of substrates by dehydrogenase hydrogen is transferred to nicotinamide 
adenine dinucleotide (NAD). The reduced NAD (NADH 2 ) transfers the hy- 
drogen normally to the electron transport chain. To get bacterial lumines- 
cence, a part of the NADH 2 is used to build reduced flavin mononucleotide 
(FMNH 2 ). FMNH 2 builds a complex with luciferase which involves the 
oxidation of a long-chain aliphatic aldehyde, developing an excited energy 
state. The complex decomposes and emits a photon. The oxidation prod- 
ucts FMN and the long chain fatty acid are reduced in the next reaction 
cycle by NADPH 2 . 

FMNH 2 + RCHO + 2 -> Luciferase -> FMN + RCOOH + H 2 + hv 

This luminescence is inhibited in the presence of hazardous substances. 
Since it is dependent on reduction equivalents, the luminescence inhibitory 
test is a physiological test belonging to the electron-transport-chain-activi- 
ty group. 

■ Procedure 

Equipment, reagents, sample preparation, procedure, and calculations are 
described in detail in ISO 1 1348 (1998). 

18.3.2 

Desmodesmus subspicatus Growth-Inhibition Assay 

■ Introduction 

Objectives. This fresh water algal growth inhibition assay is performed 
according to the standard ISO 8692 (1989). It is applicable both for the 



332 A. Eisentraeger et al. 

characterization of chemicals and aquatic environmental samples. While 
the standard allows the testing with two strains (Desmodesmus subspica- 
tus, formerly Scenedesmus subspicatus, and Selenastrum capricornutum), 
the strategy for soil characterization presented here has been set up and 
validated using the strain D. subspicatus. The algal growth inhibition test 
complements the acute bacterial luminescence test with V.fischeri. 

Principle. The growth of D. subspicatus in batch cultivation in a defined 
medium over 72 ± 2 h is quantified both in the presence and the absence 
of a sample. The cell density is measured at least every 24 h using direct 
methods like cell counting or indirect methods correlating with the di- 
rect methods, such as in vivo chlorophyll fluorescence measurement. The 
inhibition is measured as a reduction in growth rate. 

Theory. D. subspicatus is a fresh water algae that can be easily cultivated 
under defined conditions at 23 ± 2 °C with a light intensity in the range of 
35 x 10 18 to70 x 10 18 photons/m 2 /s. Since it is based on growth inhibition, 
all specific or nonspecific toxic effects relevant to reproduction of these 
algae are assessed with this test system. 

■ Procedure 

Equipment, reagents, sample preparation, procedure, and calculations are 
described in detail in ISO 8692 (1989). 

18.4 

Water-Extractable Genotoxicity 

18.4.1 

The umu Test 

■ Introduction 

Objectives. The umu test is a short-term genotoxicity assay carried out on 
microplates within less than 8 h. It is standardized for the examination of 
water and waste water (ISO 13829 2000). The water-extractable potential of 
soil samples is assessed by testing the water extract and (if the water extract 
is not genotoxic) the 15-fold concentrated water extract. The results give 
hints as to whether genotoxic substances might migrate to the groundwater. 
The umu test was chosen since it is widely applied for the examination of 
aquatic environmental samples and since both costs and time needed are 
reasonable. The procedure has been optimized and validated by charac- 
terizing large numbers of contaminated and uncontaminated soil samples 
(Ehrlichmann et al. 2000; Rila et al. 2002; Rila and Eisentraeger 2003). 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 333 

Principle. The bioassay is performed with the genetically engineered bac- 
terium Salmonella choleraesuis subsp. choleraesuis TA1535/pSK1002 (for- 
merly Salmonella typhimurium). This strain is exposed to different con- 
centrations of the samples. Different kinds of genotoxic substances can be 
detected using this test since the strain responds with different types of 
genotoxic lesions, depending on the toxin. 

Theory. The test is based on the capability of genotoxic agents to in- 
duce the umuC gene which is a part of the SOS repair system in re- 
sponse to genotoxic substances. The umuC gene is fused with the lacZ 
gene for /J-galactosidase activity. The /?-galactosidase converts ONPG (o- 
nitrophenol-/?-D-galactopyranoside) to galactose, and the yellow substance 
o-nitrophenol is quantified photometrically at 420 ± 20 nm. The tests are 
preformed both with and without metabolic activation by S9-mixture (liver 
enzymes). Cytotoxic characteristics of the samples are quantified photo- 
metrically in parallel. 

■ Procedure 

Equipment, reagents, sample preparation, procedure, and calculations are 
described in detail in ISO 8692 (1989). 

18.4.2 

Salmonella/Microsome Assay (Ames Test) 

■ Introduction 

Objectives. The Salmonella/microsome assay (Ames test) is a bacterial mu- 
tagenicity assay that is standardized according to DIN 38415 T4 (1999) for 
the determination of the genotoxic potential of water and waste water 
(Ames et al. 1975). In the strategy presented here, it is recommended if the 
umu test is negative and if there are strong hints from chemical analysis or 
site history that mutagenic compounds are present. Thus it complements 
the umu test in some cases. 

This method includes sterile filtration of the aquatic sample prior to the 
test. Due to this filtration, solid particles will be separated from the test 
sample. It may be possible that genotoxic substances are adsorbed by these 
particles. If so, they will not be detected. 

Principle. The bacterial strains Salmonella typhimurium TA 100 and TA 98 
should be used. The possible mutagenic activity of the sample is detected by 
comparing, for the bacterial strain and its activation condition, the number 
of mutant colonies on plates treated with the negative control and on plates 
treated with undiluted and diluted test samples. 



334 A. Eisentraeger et al. 

The bacteria will be exposed under defined conditions to various doses of 
the test sample and incubated for 48-72 h at 37 ± 1 °C. Under this exposure, 
genotoxic agents contained in water or waste water may be able to induce 
mutations in one or both marker genes (hisG46 for TA 100 and hisD3052 
for TA 98) in correlation with the dosage. Such induction of mutations will 
cause a dose-related increase of the numbers of mutant colonies of one or 
both strains to a biologically relevant degree above that in the control. 

Theory. Bacteria that are not able to synthesize histidine are exposed to 
mutagenous substances inducing a reversion to the wild type growing in the 
absence of histidine. Histidine auxotrophy is caused by different mutations 
in the histidine operon: S. typhimurium TA 98 contains the frameshift 
mutation hisD3052 reverting to histidine independency by addition or 
loss of base pairs. S. typhimurium TA 100 bears the base pair substitution 
hisG46 which can be reverted via base pair substitutions (transition or 
transversion). 

The tester strains are deep rough enabling larger molecules also to pen- 
etrate the bacterial cell wall and produce mutations (rfa mutation). The 
excision repair system is disabled (Z\uvrB), increasing the sensitivity by 
reducing the capability to repair DNA damage. Furthermore, they contain 
the plasmid pKMlOl coding for an ampicillin resistance. 

■ Procedure 

Equipment, reagents, sample preparation, procedure, and calculations are 
described in detail in DIN 38415 T4 (1999). An ISO standard is in prepara- 
tion. 

18.5 

Habitat Function: 

Soil/Microorganisms, Soil/Soil Fauna, Soil/Higher Plants 

18.5.1 

Respiration Curve Test 

■ Introduction 

Objectives. The determination of respiration curves provides information 
on the microbial biomass in soils and its activity. The method is suitable 
for monitoring soil quality and evaluating the ecotoxicological potential of 
soils. It can be used for soils sampled in the field or during remediation 
processes. The method is also suitable for soils that are contaminated 
experimentally either in the field or in the laboratory (chemical testing). 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 335 

Principle. The C0 2 production or 2 consumption (respiration rate) from 
unamended soils as well as the decomposition of an easily biodegradable 
substrate (glucose + ammonium + phosphate) is monitored regularly (e.g., 
every hour). From the C0 2 -production or 2 -consumption data the dif- 
ferent microbial parameters, such as basal respiration, substrate-induced 
respiration, lag time, are calculated. 

Theory. Basal respiration and substrate-induced respiration (SIR) are wide- 
ly used physiological methods for the characterization of soil microbial 
activity and biomass. Basal respiration gives information on the actual 
state of microbial activity in the soil. After addition of an easily biodegrad- 
able carbon source respiration activity increases. At the time of substrate 
addition the activity can be described by 

SIR = r + K 

where r is the initial respiration rate of growing microorganisms. 

In the course of an incubation period the respiration rate increases and 
can be described by 

dp/dt = rept + K 

This equation is based on the assumption that the increase of the respi- 
ration rate dp/dt after substrate addition in the SIR method represents 
the sum of the respiration rates of growing (rept) and non-growing (K) 
microorganisms (Stenstrom et al. 1998). 

The microbial respiration activity is affected by several parameters. Wa- 
ter content, temperature (Blagodatskaya et al. 1996), the quality of the soil 
organic matter (Wander 2004), as well as contaminants (e.g., Blagodatskaya 
and Anan'eva 1996; Kandeler et al. 1996) show an influence. 

■ Procedure 

Sample preparation, equipment, reagents, procedure, and calculations are 
described in detail in ISO 17155 (2002). A prerequisite is equipment that 
allows the determination of C0 2 release or 2 uptake at short time in- 
tervals. Basal respiration is measured first. The respiration rates should 
be measured until constant rates are obtained. After measuring the basal 
respiration, a defined substrate mixture containing glucose, potassium di- 
hydrogen phosphate, and diammonium sulfate is added. The mixture is 
made up of: 80 g glucose, 13 g KH 2 P0 4) and 2 g (NH 4 ) 2 S0 4 . In testing, 0.2 g 
mixture is used per gram of soil in which at least 1 g organic matter is 
found in 100 g soil dry mass. The measurement of C0 2 evolution or 2 
consumption has to be continued until the respiration curve reaches its 
peak and the respiration rates are declining. 



336 A. Eisentraeger et al. 

The ecotoxicological potential of soils is described by several parameters: 

• Respiratory activation quotient: basal respiration rate divided by sub- 
strate-induced respiration rate (Qr = Rb/Rs) 

• Lag time (ti ag ): the time from addition of a growth substrate until ex- 
ponential growth starts, - a reflection of the vitality of the growing 
organisms 

• Time to the peak maximum (t pea kmax): the time from addition of growth 
substrate to the maximum respiration rate - another reflection of the 
vitality of the growing organisms 

According to the guideline, Q R > 0.3, ti ag > 20 h, and t pea kmax > 50h 
indicate polluted materials. 

■ Notes and Points to Watch 

• Increased respiratory activation quotients may occur for two reasons. 
On one hand, they are an indicator of bioavailable carbon sources. These 
may be of biological origin, as for example compost, or biodegradable 
organic contaminants (e.g., mineral oil, anthracene oil, phenanthrene) 
that have the same effect (Hund and Schenk 1994). Sufficient amounts 
of biodegradable carbon sources always result in increased respiration 
activities when a sufficient amount of further nutrients (e.g., nitrogen, 
phosphate) is available. On the other hand increased Q R s may be an 
indicator of contaminants that are not biodegradable, e.g., heavy metals 
(Nordgren et al. 1988). Up to now, it is not known how to distinguish 
which parameters are responsible for a stress-induced respiration caus- 
ing increased quotients. 

• It has to be considered for the assessment that increased values indi- 
cate amended/contaminated soils, whereas not all contaminated soils 
show higher values. Accordingly, it cannot be concluded that the habitat 
function of a soil is intact when the respiration values are in a normal 
range. 

• In the literature, the derivation of a metabolic quotient (basal respira- 
tion divided by microbial biomass) as an indicator for an ecosystem is 
described (Insam and Domsch 1988; Anderson and Domsch 1990). In 
soils with a recent input of easily biodegradable substrates, mainly r- 
strategists occur. They usually respire more C0 2 per unit degradable C 
than k-strategists, which prevail in soils that have not received fresh or- 
ganic matter and have evolved a more complex detritus food web (Insam 
1990). Since the substrate-induced respiration can be used to calculate 
the microbial biomass, it could be concluded that the metabolic quotient 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 337 

and the respiration activation quotient are comparable. In this context it 
should be noted that the estimation of the microbial biomass by Ander- 
son and Domsch (1978) is based on a linear regression between SIR and 
the microbial biomass according to the fumigation- incubation method. 
The conversion factor was elaborated on the basis of a range of soils. 
However, in other soils the population may differ from the originally 
investigated soils (e.g., forest soils vs. contaminated soils) and different 
conversion factors may be necessary (Hintze et al. 1994). One should, 
therefore, avoid calculating the microbial biomass of soils on the basis 
of the substrate-induced respiration for which the conversion factor is 
unknown. 



18.5.2 

Ammonium Oxidation Test 

■ Introduction 

Objectives. This test is a rapid procedure for determining the potential 
rate of ammonium oxidation in soils. The method is suitable for all soils 
containing a population of nitrifying organisms. It can be used as a rapid 
screening test for monitoring the quality of soils and wastes, and it is 
suitable for testing the effects of cultivation methods, chemical substances, 
and pollution in soils. 

Principle. Ammonium oxidation, the first step in autotrophic nitrification 
in soil, is used to assess the potential activity of microbial nitrifying pop- 
ulations. Autotrophic ammonium-oxidizing bacteria are exposed to am- 
monium sulfate in a soil slurry. Oxidation of the nitrite formed by nitrite- 
oxidizing bacteria in the slurry is inhibited by the addition of sodium 
chlorate. The subsequent accumulation of nitrite is measured over a 6-h 
incubation period and is taken as an estimate of the potential activity of 
ammonium oxidizing bacteria. For the assessment of soil quality the nitri- 
fication activity in a test soil, in a control soil, and in a mixture of both soils 
is determined. 

Theory. In soils with pH > 5. 5 nitrification is performed by chemoau- 
totrophic nitrifiers (Focht and Verstraete 1977). The procedure consists of 
two steps. Ammonium is oxidized to nitrite by one group of nitrifiers, while 
nitrite is oxidized to nitrate by a second group. Since nitrite is oxidized as 
it is produced, the rate at which ammonium is oxidized is equal to that at 
which nitrite plus nitrate accumulate. To avoid the application of two meth- 
ods - one for the determination of nitrite and one for determining nitrate - 
a procedure was developed to completely and specifically block the oxida- 
tion of nitrite. With this method it is possible to get information on the 



338 A. Eisentraeger et al. 

nitrification process by using only one analytical method, since the rate at 
which nitrite alone accumulates equals the rate of ammonium oxidation. In 
soils with a high background of nitrate this method is much more sensitive, 
since nitrite normally is undetectable at the beginning of the incubation. 
A prerequisite for a correct measurement is (1) that the inhibitor does not 
inhibit ammonium oxidation, and (2) that the inhibitor completely blocks 
nitrite oxidation. Chlorate has proved to be an appropriate inhibitor. At 
suitable concentrations an inhibition of ammonium oxidation seems to be 
negligible. Although, in some cases, the inhibition of nitrite oxidation can 
be incomplete, this does not seem to be a real problem. It is negligible when 
Vmax f° r nitrite oxidation is lower than the rate of ammonium oxidation. It 
might be a problem, if V max is larger. Since chlorate mainly influence the K m 
of the reaction, the initial rate of the reaction is the best estimate of the am- 
monium oxidation rate. Leakage will be lowest at low nitrite concentrations 
(Belser and Mays 1980). 

The results present a potential activity, since several test parameters are 
different from natural conditions: Ammonium is added in surplus, aeration 
is probably more intensive by shaking in the laboratory than under field 
conditions, and the incubation temperature of 25 °C usually far exceeds 
real soil conditions. 

Several methods exist to get information on nitrification in soil. Some 
of these are characterized by incubation periods of several weeks (e.g., ISO 
14238 1997). For soil assessments the determination of the ammonium 
oxidation activity was selected since this procedure has several advantages, 
especially for investigation of contaminated soils and for soil remediation 
procedures. These applications frequently require results within a short 
period of time, as they contribute to decisions whether a soil has to be 
remediated, whether a remediation has to be continued, or whether the 
habitat function of the soil (at least with respect to microorganisms) is intact 
so that the soil can leave the remediation plant. This is important in avoiding 
unneeded and expensive retention of soil in the remediation plants. As the 
potential ammonium oxidation method yields results in a short period 
of time, and furthermore is suitable for soils with high nitrate contents 
(during bioremediation nitrogen has to be added to achieve degradation 
of contaminants), this method was selected for the ecotoxicological soil 
assessment. 



■ Procedure 

Sample preparation, equipment, reagents, procedure, and calculations are 
described in detail in ISO 15685 (2004). For soil assessments three different 
test designs are applied: 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 339 

1. Test soil 

2. Reference soil (uncontaminated soil with a nitrification activity of about 
200-800 ng N/g dry mass of soil/h) 

3. Mixture of test soil and reference soil (1:1 with regard to soil dry mass) 

The soils are adjusted to 60% of WHC max and incubated for 2 days at 20 ° C. 
The mixture is prepared immediately before testing. The mixture and the 
two soils are incubated again for 1 day at 20 °C, after which the nitrification 
activity is determined. Soil samples are mixed with test medium containing 
phosphate, sodium chlorate and diammonium sulfate. The slurries are 
incubated for 6h at 25 ± 2°C on an orbital shaking incubator (about 
175 rpm). 2-mL samples are taken after 2 and 6 h, and the nitrite content is 
determined. The mentioned time interval is a recommendation. 

■ Calculation 

The rate of ammonium oxidation (ng NO2 -N /g dry mass of soil/h) is 
calculated from the difference of NO2 -N concentrations at the different 
measuring times. The following formula is applied for the assessment of 
the test soil: 

M m + SD m < 0. 9 x (M c - M P )/2 (18.1) 

M m mean ammonium oxidation activity in soil mixture 

SD m standard deviation of ammonium activity in replicate test vessels with 
soil mixture 

M c mean ammonium oxidation activity in control soil 

M P mean ammonium oxidation activity in polluted soil 

The polluted soil is considered to be toxic if the mixture has an am- 
monium oxidation activity significantly slower than 90% of the calculated 
mean activity of the two single soils. 

■ Notes and Points to Watch 

• The suitability of storing soil samples at -20 °C is discussed controver- 
sially. The investigation of 12 soils differing in their physico-chemical 
properties has revealed that storage at -20 °C for 13 months does not 
affect the nitrifiers in annually frozen soils in any decisive way (Sten- 
berg et al. 1998). As the procedure, however, does not seem to be suitable 
for every soil, in the guideline ISO 15685 (2004) storage at -20 °C is not 
generally recommended. The different results found in the literature on 
the effects of freezing as a storage method can be explained in a number 



340 A. Eisentraeger et al. 

of ways: The populations in soils annually subjected to several freeze and 
thaw cycles seem to be adapted and more resistant to freezing than the 
microflora in soils where freeze and thaw cycles are not a regular occur- 
rence. Furthermore, the growth status of the microorganisms at the time 
of sampling may play a role. Active cells seem to be more sensitive to 
freezing and thawing than less active cells. Therefore, samples collected 
shortly after managing processes such as fertilizing or tilling may show 
cell depletion. Furthermore, the selected procedure of freezing and thaw- 
ing may influence the results. Slow rates of temperature change seem to 
result in greater microbial losses. Storage in small portions and rapid 
temperature flux maybe preferable (Stenberg et al. 1998). In conclusion, 
soils should only be stored if the effect is known and acceptable. 

18.5.3 

Combined Earthworm Mortality/Reproduction Test 

■ Introduction 

Objectives. The determination of the survival and the reproductive success 
of earthworms as representatives of soil macrofauna provide information 
on these saprophagous soft-bodied invertebrates that in many soils play an 
important role as ecosystem engineers. The method is suitable for moni- 
toring soil quality and the evaluation of the ecotoxicological potential of 
soils. It can be used for soils sampled in the field or during remediation pro- 
cesses. Furthermore the method is suitable for soils that are contaminated 
experimentally in the field or in the laboratory (e.g., chemical testing, in 
particular pesticide testing). 

Principle. Adult earthworms are either exposed to potentially contami- 
nated soil samples or to a range of concentrations of a test substance mixed 
in an artificial or natural control soil. The mortality and the biomass of the 
adult worms are measured after 28 days. The effect on the reproduction is 
determined by counting the number of juveniles hatched from the cocoons 
after an additional period of 4 weeks. Based on these measurements, the 
ecotoxicological potential of the test soil is determined. 

Theory. Earthworms are important members of the soil community due 
to their ability to change or create their habitat through various activities, 
thus correctly considered to be "ecosystem engineers" (Lavelle et al. 1997): 

• Penetrating the soil and building burrows, as well as increasing pore 
space 

• Transporting soil and organic matter by casting 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 341 

• Comminuting organic material (including cattle feces in meadows) as 
a first step in its breakdown 

• Providing nutrients to plants (e.g., by concentrating them in burrow 
linings or by increasing the availability of nutrients like phosphorus) 

• Relocating seeds in the soil profile 

• Changing the diversity and improving the activity of the microbial com- 
munity by selective feeding and providing feces rich in nutrients 

Finally, earthworms are closely exposed to all contaminants occurring 
in the soil solution but also - by feeding - to all chemicals adsorbed to soil 
particles. 

These activities thus finally lead to an improved soil structure, i.e. to 
stabilization of soil aggregates, to increase in water infiltration (partly by 
higher water-holding capacity; Urbanek and Dolezak 1992; Edwards and 
Shipitalo 1998), often to the formation of a humic layer close to the soil 
surface (mainly in forest ecosystems; Doube and Brown 1998), and to an 
increased yield in orchards or grassland (Blakemore 1997). The activities 
described above are performed by various earthworm species to a very dif- 
ferent extent. Still, large, deep -burrowing worms like Lumbricus terrestris 
are involved in several of these activities, especially concerning soil struc- 
ture and organic matter breakdown (Swift et al. 1979). In the light of this 
knowledge, it is difficult to understand why the main earthworm species 
used in tests are the two closely related compost worms Eisenia fetida or 
Eisenia andrei. Ecologically, these species are less important than the deep- 
burrowing worms (L0kke and van Gestel 1998). On the other hand, from 
a practical point of view the compost worms are more suitable than any 
other lumbricid species because they reproduce very quickly and easily in 
the laboratory, and mass cultures can be obtained. In addition, the sensi- 
tivity of these species is in the same general order of magnitude as other 
earthworm species. In most cases the differences between species are, de- 
pending on the chemical or contaminant mixture tested, not larger than by 
a factor of 10 (Roembke 1997; Jones and Hart 1998). 

Concerning the test endpoints, the determination of mortality covers 
strong acute effects. However, from an ecological point of view such effects 
are clearly less important than long-term, chronic effects usually occurring 
at relatively low and thus more realistic concentrations (see "Notes and 
Points to Watch"). For this reason, reproduction is the test variable of 
highest relevance. 

■ Procedure 

Equipment, reagents, sample preparation, procedure, and calculation of 
the test results are described in detail in the ISO guidelines 1 1268-1 (1993) 



342 A. Eisentraeger et al. 

and 11268-2 (1998). In deviation from these guidelines in which the acute 
and chronic endpoints are determined in individual test runs, it is recom- 
mended to use a combined test method for the assessment of contaminated 
soils. For the assessment of single chemicals, separate tests should still 
be used in order to be in agreement with legal requirements concerning 
the risk assessment of chemicals (e.g., the EU guideline describing the 
registration of pesticides; European Union 1991). 

Ten adult earthworms of the species E.fetida or E. andrei per test vessel 
are exposed to a series of mixtures of the potentially contaminated test soil 
and an uncontaminated control or reference soil at 20 ± 2 °C for 4 weeks. 
If the mortality in the contaminated test soil is higher than 20%, the test is 
stopped. Otherwise, at the end of this period, the adult worms are removed 
from the vessels and the surviving animals are counted and weighed. After- 
wards, the test soil remains in the same vessels for another 4 weeks. After 
56 days the juveniles are extracted from test and control soils and counted. 
For the endpoint reproduction the data of the test soil vessels are compared 
with those from the controls. An inhibition of reproduction of 50% com- 
pared to the control is indicative of a contaminated soil sample. A soil that 
causes mortality higher than 20% is also classified as contaminated. 

■ Notes and Points to Watch 

• As already mentioned, the acute test endpoint mortality is ecologically 
not relevant due to the following reasons: Lumbricid worms die slowly 
and only at high concentrations of soil contaminants. In real field situa- 
tions (with the exception of relatively small areas like mining deposits) 
the concentrations of chemicals are low but these substances, in particu- 
lar metals, are often persistent. Such effects are much better determined 
by using chronic sensitive endpoints like reproduction. Ecologically, in 
many populations of earthworms any impact more strongly affects the 
reproductive rate than it does mortality rate. A short-term decrease in 
the number of individuals is easier to compensate than a long-term re- 
duction in the number of juveniles. For this reason, the assessment of 
the biological quality of soil should be based on the chronic endpoint 
reproduction. 

18.5.4 

Collembola Reproduction Test 

■ Introduction 

Objectives. The determination of the survival and the reproductive success 
of collembolans as representatives of soil mesofauna provides information 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 343 

on these saprophagous hard-bodied invertebrates, an important part of 
the soil food web in many soils. The method is suitable for monitoring soil 
quality and evaluating the ecotoxicological potential of soils. It can be used 
for soils sampled in the field or during remediation processes. Furthermore, 
the method is suitable for soils contaminated experimentally in the field or 
in the laboratory (e.g., chemical testing, in particular pesticide testing). 

Principle. Juvenile collembolans are either exposed to potentially contam- 
inated soil samples or to a range of concentrations of the test substance 
mixed in artificial soil. The mortality of the adult springtails as well as the 
reproduction (= number of juveniles) are measured at the end of the expo- 
sure period of 28 days. Based on these measurements, the ecotoxicological 
potential of the test soil is determined. 

Theory. The species Folsomia Candida (Collembola) is tested as a repre- 
sentative of hard-bodied soil invertebrates, in particular arthropods (Ac- 
hazi et al. 2000). These organisms, mainly consisting of springtails (Collem- 
bola) and mites (Acari), are among the most numerous invertebrates in 
a wide range of soil types, especially of the Northern hemisphere. Due 
to their high numbers they are an important part of the soil food web 
(Weigmann 1993). In addition, the springtails control by their feeding ac- 
tivity the population cycles of microorganisms, which in turn are extremely 
important as mineralizers of organic matter (Swift et al. 1979). To a lesser 
extent, springtails can also influence the numbers of nematodes (Hopkin 
1997). Finally, they are exposed to contaminants via pore water and air 
space. 

The species F. Candida is distributed worldwide (mainly by anthro- 
pogenic activities). It prefers soils with an elevated content of organic 
matter but is not only a compost inhabitant (e.g., it occurs in comparatively 
low numbers in agricultural soils; Petersen 1994; Hopkin 1997). Its use is 
criticized for the same reasons discussed for compost worms. However, 
the response is similar: F. Candida is easily cultured and its sensitivity, as 
far as known, is not considerably different from other collembolans (Ac- 
hazi et al. 2000). As in the case of earthworms, the endpoint reproduction 
is ecologically highly important (see Sect. 18.5.5). 

■ Procedure 

Equipment, reagents, sample preparation, procedure, and calculation of the 
test results are described in detail in the ISO guideline 11267 (1999). Ten 
juvenile springtails of the species F. Candida per test vessel are exposed to 
a potentially contaminated soil sample or a series of mixtures between the 
test soil and an uncontaminated control or reference soil (plus a control) at 
20±2 ° C for 4 weeks. At the end of this period, the collembolans are removed 



344 A. Eisentraeger et al. 

from the vessels and the surviving animals are counted (juveniles and 
adults separately) by using photographs or an automatic image processing 
system. For the endpoint reproduction, the data from the test soil vessels 
are compared with the controls. An inhibition of reproduction of 50% 
compared to the control is indicative of a contaminated soil sample. 

■ Notes and Points to Watch 

• The common test species F. Candida is difficult to distinguish from other 
species of the same genus, in particular F. fimetaria (Wiles and Krogh 
1998). This species has also been proposed for ecotoxicological testing, 
but it reproduces sexually and is, as such, more difficult to handle. Due to 
such practical problems and since it is not known whether the two species 
are equally sensitive to chemicals, any mixing of them must be carefully 
avoided. In cases of doubt a taxonomist specialized in collembolans 
should be consulted. 



18.5.5 

Plant Growth Test 

■ Introduction 

Objectives. The determination of the emergence and growth of different 
plant species allows assessment of the quality of a certain soil as a habitat 
for terrestrial primary producers (i.e., in terms of nutrient cycling, the basis 
of the whole ecosystem). The method is suitable for monitoring soil quality 
and evaluating the ecotoxicological potential of soils. It can be used for 
soils sampled in the field or during remediation processes. Furthermore, 
the method is suitable for soils that are contaminated experimentally in the 
field or in the laboratory (chemical testing, in particular pesticide testing). 

Principle. This phytotoxicity test is based on the emergence and early 
growth response of a variety of terrestrial plant species to potentially 
contaminated soil. Seeds of selected species of plants are planted in pots 
containing the test soil and in control pots. They are kept under growing 
conditions for the chosen plants and the emergence and mass of the test 
plants are compared against those of control plants. 

Theory. The importance of plants as the basis of ecosystem performance, 
but also for the production of food and forage, cannot be overestimated 
(Riepert et al. 2000). In 1984, plants were added to the list of terrestrial 
test species by the OECD. These selected species still represent agricultural 
plants, while wild herbs, trees, etc., are usually not tested (Boutin et al. 1 995). 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 345 

For the testing of chemicals, often two exposure pathways are distinguished: 
airborne via aboveground plant parts (e.g., after the spraying of pesticides) 
or via soil mixtures. Obviously, in the case of contaminated soil only the 
latter test version is used. 

Concerning the measurement endpoints, the fresh biomass of the above- 
ground parts has been selected due to the practicability of evaluating it and 
its high sensitivity. However, one must be aware that this selection has been 
done for an acute test with a duration of 1 4 days. Further research will clarify 
whether long-lasting chronic tests (e.g., using the endpoint reproduction) 
will be more sensitive (ISO 22030 2004). 

■ Procedure 

Equipment, reagents, sample preparation, procedure, and calculation of 
the test results are described in detail in the ISO guideline 11269-2 (1995). 
In supplementing the guideline the test was changed in two ways: 

1 . In addition to the pure test soils, mixtures of the potentially contaminated 
soils with a suitable control or reference soil are made in a ratio of 
50:50. 

2. While the ISO lists 15 potential test species, it is recommended to use 
only the monocotyledonous species Avena sativa (oat) and one of the two 
named dicotyledonous species, either Brassica rapa (turnip) or Lepidum 
sativum (cress), for soil quality assessment. Each treatment is tested in 
four replicates (10 seeds per replicate (= test vessel)). Watering is done 
by using a semi-automated wick method (Stalder and Pestemer 1980). 
After emergence, the seedlings are thinned to a final number of five per 
vessel. Fourteen days later the aboveground parts of the plants (fresh 
mass) are harvested and weighed. 

■ Evaluation 

The evaluation is done according to the following formula (Winkel and 
Wilke 2000): 

M g + SD Mg < 0. 9 x M b (18.2) 

M g Biomass measured in the vessels with the 50:50 mixture of test 

and control soil 

SD Mg Standard deviation of the 50:50 mixture between test and control 

soil 



346 A. Eisentraeger et al. 

0. 9 x Mb The calculated mean between the test and the control soil (bio- 

mass t est soil + biomass C ontroisoii) divided by 2 minus a tolerance 
value of 10%. 

A soil is classified as toxic if the biomass measured in the vessels with 
the 50:50 mixture of test and control soil is > 10% lower than the mean 
biomass determined in the test and control soils. 



Notes and Points to Watch 

In addition to storage problems already mentioned in the context of other 
terrestrial tests, it must be pointed out that in the case of plant testing the 
amount of plant- available nitrogen is very important for the growth of 
the test organisms, including the controls. If the plants grow badly in the 
controls it is difficult to identify effects occurring in the test vessels with 
test soils. For this reason, Riepert and Felgentreu (2000) recommended 
to avoid the use of fresh field soils because they don't contain enough 
available nitrogen due to high microbial activity. In order to solve this 
general problem fertilizer could be added to the water reservoirs used in 
the plant tests. Since all plants (both in the test as well as in the control 
vessels) are on the same nutrient level any effect caused by nitrogen 
availability would be eliminated. However, one must be cautious since 
some soils might be already so rich in nutrients that over-fertilization 
could occur. 

Another problem in testing potentially contaminated soils with plants 
is the fact that structural properties of the soil can affect the plants 
too. If the habitat function of the soil has to be assessed in general, the 
distinction between chemical and physical properties is not necessary. 
However, there are many field soils which are not suitable for the growth 
of crop species (e.g., acid soils). In order to avoid false positive results, 
the ecological requirements of the common test species (oat, turnip) 
are currently being studied (Jessen-Hesse et al. 2003). These data will 
allow the determination of which soils can be tested with the current test 
species and which cannot. 



18.5.6 

Test Performance for the Derivation of Threshold Values 

■ Introduction 

Objectives. The described terrestrial ecotoxicological tests are also suitable 
for the derivation of threshold values to protect the habitat function of 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 347 

soils for soil organisms. The protection of this soil function is required in 
the German Soil Protection Act (BBodSchG 1998). The threshold values 
indicate the contamination pathway soil to soil organisms. 

Principle. Soils are contaminated experimentally and the biological effect 
is investigated (chemical testing). Several concentrations are tested and the 
ecotoxicological potential is determined. Based on these measurements 
LC 50 (lethal concentration) or EC 50 (effective concentration) values for the 
different endpoints are calculated, using appropriate statistical methods. 

Theory. Ecotoxicological tests provide information on the toxicity of pri- 
ority contaminants. For the derivation of trigger values it has to be kept 
in mind that only a limited number of species and organisms have been 
tested. To protect the "whole" ecosystem, extrapolation methods have to 
be applied. Depending on the amount of available data the extrapolation 
method DIBAEX (distribution based extrapolation; Wagner and L0kke 
1991) or FAME (factorial application method; European Union 1996) may 
be suitable. For the derivation of trigger values concerning the pathway 
soil to soil organisms, this procedure was successfully applied in Germany 
(Wilke et al. 2001; Wilke et al. 2004). In Germany trigger values are those 
which, if exceeded, indicate a harmful soil change or site contamination. 
If such cases occur, investigations of the site have to be performed. Since 
they indicate a potential effect, EC 50 and LC 50 values instead of NOEC 
(no-observed-effect concentration) or LOEC (lowest-observed-effect con- 
centration) values are applied for the derivation. 

■ Procedure 

Chemical testing is described in detail in the different guidelines (mainly 
from OECD) mentioned in the pertinent sections. 

■ Notes and Points to Watch 

• Control soils have to be selected carefully (ISO 15799 2004). For the 
derivation of trigger values, natural soils are recommended, but at least 
a sandy soil with low sorption capacity should be used. For higher 
environmental relevance, loamy and silty soils should be employed. If 
artificial soil is used and if the test chemical has a high logK ow value 
(octanol-water partitioning coefficient; e.g., > 2; European Plant Protec- 
tion Organization 2003) this test substrate should contain only 5% peat 
instead of 10% in order to test a more field-relevant situation concerning 
the bioavailability of the test substance. 



348 



A. Eisentraeger et al. 



18.6 

Combined Performance of Bioassays 

and Assessment of the Results 

18.6.1 

Water-Extractable Ecotoxic Potential 

The procedure proposed here, and based on only two bioassays, is a qual- 
itative one that offers a way to quickly obtain results and keep costs down 
(Fig. 18.2). Dilution values are defined to indicate ecotoxicological poten- 
tial if exceeded. The water extracts should be diluted using a factor of 2, 
and lowest ineffective dilution values (LID) should be assessed. The LID 
is defined as the lowest dilution with less than 20% inhibition in the lu- 
minescence algae test. For the qualitative evaluation it is not necessary to 
determine EC values by data transformation from dose response curves. Of 
course, it might be useful to determine EC values if toxic potentials of soil 
samples (e.g., from the same site during remediation) have to be compared. 
The V. fischeri luminescence inhibition assay (ISO 11348 1998; Sect. 
18.3.1) should be performed at first. If the LID value exceeds 8, a risk of 
pollutant leakage exists and it is recommended that the remediated soils 
should not be incorporated at unsealed sites. If the luminescence inhibi- 
tion assay is negative, the 72 h algae growth inhibition assay (ISO 8692 



Vibrio fischeri luminescence 

inhibition test 

(ISO 11348) 

or 

Desmodesmus subspicatus 

growth inhibition test (ISO 8692) 



LID>8 



LID>4 



^> 



Risk of pollutant leakage is 
given 

No incorporation of 

remediated soil at unsealed 

sites 



Vibrio fischeri luminescence 

inhibition test 

(ISO 11348) 

and 

Desmodesmus subspicatus 

growth inhibition test (ISO 8692) 



LID<8 



LID<4 



■=> 



Risk of pollutant leakage is 
low 



Fig. 18.2. Procedure proposed for the assessment of the water extractable ecotoxicological 
potential of soils and soil materials. The assessment based on LID values is allowed if 
(1) a dose response relationship is obtained, or (2) nearly 100% inhibition is obtained in 
several tested dilutions, or (3) no significant inhibition is obtained in the dilutions up to the 
threshold value. (Maxam et al. 2000; Pfeifer et al. 2000; Dechema 2001; Rila and Eisentraeger 
2003; Eisentraeger et al. 2004) 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 349 

1989; Sect. 18.6.2) should be performed additionally. The risk of pollutant 
leakage is low if this LID value is < 4 (or the LID value of the lumines- 
cence inhibition assay is < 8). These threshold values are derived from 
the experiences gained during the earlier-mentioned research projects 
(Rila and Eisentraeger 2003) and from the results of a ring test (Hund- 
Rinke et al. 2002a, b). Low inhibitions are obtained with uncontaminated 
soil samples such as the natural standard soils LUFA 2.1, 2.2 and 2.3 (land- 
wirtschaftliche Untersuchungs- und Forschungsanstalt, Speyer, Germany). 
This "background toxicity" might be caused by humic substances. 

Ecotoxic effects of a wide range of water-extractable contaminants can 
be detected by using these two test systems. In a round robin test eight 
contaminated soils were investigated using four aquatic test systems (lumi- 
nescence and growth test with V.fischeri, tests with algae and daphnids). It 
was shown that daphnids are mostly less sensitive than the tests with algae 
and the luminescence test with V.fischeri. The daphnids test was more sensi- 
tive, however, for soils contaminated with heavy metals (Hund-Rinke et al. 
2002c). As heavy metals are routinely measured by chemical analyses, it 
was decided to exclude the test with daphnids from the base set of aquatic 
ecotoxicological test systems for soil assessment. The approach presented 
here is cost effective: No range-finding test has to be carried out and the 
algae growth inhibition test can be performed in microplates, so long as 
the validity criteria of ISO 8692 are fulfilled (Eisentraeger et al. 2003). If 
other or further testing is regarded as necessary, ecological relevance and 
practicability should be considered. 



18.6.2 

Water-Extractable Genotoxicity 

Cost effectiveness and speed are also major aspects of the assessment 
scheme for water-extractable genotoxic potential. It should thus be noted 
that this is a screening method that cannot be used to identify clastogenic 
substances, but is able to roughly estimate whether genotoxic compounds 
can be mobilized by water. The procedure is mainly based on the assessment 
of the genotoxic potential of water extracts using the umu test according to 
ISO 13829 (2000; Sect. 18.4.1). The umu test can be performed in less than 
a day with and without metabolic activation. The Salmonella/microsome 
test (Ames test) according to DIN 38415 T4 (Sect. 18.4.2) should be carried 
out additionally if the umu test is negative and if there are strong hints from 
chemical analysis or site history that mutagenic compounds are present. 

In the first step of the procedure (Fig. 18.3) the same water extract is 
tested as used for the assessment of the water-extractable ecotoxicological 
potential. If there is a genotoxic effect in the umu test, with or without 



350 



A. Eisentraeger et al. 





First step: 


umu-test 
optional: 
Ames-test 


D Li >3 
LID>6 




umu-test 
optional: 
Ames-test 


D LI =1.5 
LID = 3 


Second step: 


Investigation of coi 


umu-test 
optional: 
Ames-test 


D Li >3 
LID>6 




umu-test 
optional: 
Ames-test 


Dii=1.5 
LID = 3 



Investigation of the soil elutriates 



genotoxic 



^> 



acute danger for path soil- 
groundwater 



not 
genotoxic 



^> 



Risk of acute danger for 

path soil-groundwater 

is low 



genotoxic 



^> 



acute danger for path 
soil-groundwater 



not 
genotoxic 



^> 



no danger for path 
soil-groundwater 



Fig. 18.3. Assessment of the water-extractable genotoxic potential of soils and soil materials 
using the umu test according to ISO 13829. The Salmonella/microsome test (Ames test) 
according to DIN 38415 T4 should be carried out if the umu test is negative and there are 
strong hints from chemical analysis or site history that mutagenic compounds are present. 
(Eisentraeger et al. 2000; Dechema 2001; Eisentraeger et al. 2001; Rila and Eisentraeger 2003; 
Eisentraeger et al. 2004; modified according to Ehrlichmann et al. 2000) 



metabolic activation, a high risk of transfer of genotoxic substances from 
soil to the ground water exists. If there is no genotoxic effect, the water 
extract should be concentrated by a (low) factor of 15 using Serdolit PAD- 
1 resin. During the ring test mentioned above (Hund-Rinke et al. 2002a) 
the water extracts were concentrated by a factor of 30, as performed by 
Ehrlichmann et al. (2000). The factor was reduced to 15 on the basis of 
results obtained during this test and further studies (Rila and Eisentraeger 
2003), since several obviously uncontaminated soil samples (e.g., LUFA 2.1 
and LUFA 2.2) tested positive after 30-fold concentration. 

18.6.3 

Assessment of the Habitat Function 



Criteria for the combined assessment of the pathways from soil to soil 
microorganisms, fauna, and higher plants were elaborated (Fig. 18.4). The 



18 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 



351 











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o 




go O 

CD 5-i 


"3 




m 






PQ 




A 




d o 


X 




d 




ft ° 


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352 A. Eisentraeger et al. 

test design was approved in a round robin test. For special requirements 
it is possible to complement the basic test set by further tests. Test results 
are interpreted using different strategies selected, depending on the test 
system employed. 

Soil Microflora - Respiration Activity. The respiration activity is assessed by 
the ratio basal respiration:SIR, and by considering the 2 uptake or C0 2 
production over time. 

Soil Microflora - Ammonium Oxidation Activity. The nitrification activity of the 
test soil is assessed by comparison with a control soil and a 1:1 mixture of 
test soil and control soil. If the nitrification activity in the mixture is below 
90% of the mean value of the activity in the test and control soil, the habitat 
function is assessed as "disturbed" for this criterion. 

Soil Fauna. Regarding soil fauna, a minimal habitat function is demanded. 
The assessment is based on the comparison between the mortality rate 
and the reproduction in the test soil and in the control. The habitat func- 
tion is considered disturbed if the mortality rate surpasses 20% and the 
reproduction rate falls below 50% compared to the control. 

Soil Flora. To evaluate potential effects on the soil flora two test strategies 
have been elaborated. For both strategies a control soil is needed. The first 
strategy directly compares the biomass production in the test soil and in 
the control soil. A second possibility is to compare the biomass production 
in (1) the test soil, (2) a control soil, and (3) a 1:1 mixture of the test and 
control soils. A biomass determined to be less than 70% in the test soil as 
compared to the control or less than 90% in comparison to the mean value 
of the mixed test and control soils is regarded as insufficient and the sample 
is assessed "toxic". 

Preferably, a control soil from the site should have the same physico- 
chemical soil properties as the contaminated soil but no contamination. 
However, in many cases such a soil is not available and it is then recom- 
mended to use a sandy soil (e.g., LUFA standard soil 2.2) to avoid a high 
sorption of contaminants (for more details see ISO 15799 2003). In cases 
where the geographical or pedological typicality of the selected soil is 
important, approaches like the EURO Soil concept (Kuhnt and Muntau 
1992) or the German Refesol proposal can help to find appropriate control 
soils. 

The terrestrial tests were selected to give information on the habitat 
function of the soil. If the habitat function of a soil is reduced, this may 
result from anthropogenic contaminants (e.g., heavy metals, PAHs, TNT), 
a high salt content caused by the addition of large amounts of organic 
material (e.g., compost), or alow pH. Therefore, expert knowledge is needed 
to decide whether a test is suitable for a specific soil or soil material. 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 353 

Moreover, results indicating a toxic potential have to be critically examined 
with respect to further decisions regarding the use of the test material. 
If there seems to be a need to replace a test or to perform further tests, 
ecological relevance and practicability should be considered. Under certain 
circumstances, field monitoring approaches at the assessment site may be 
appropriate (Roembke and Notenboom 2002). 

18.6.4 

Overall Assessment - Combined Strategy 

In Fig. 18.5, a stepwise procedure for the combined evaluation of remediated 
soil samples is given as an example for the cost effective application of these 
bioassays. 

1. In the first step it is determined by chemical analyses whether thresh- 
old values for single contaminants are exceeded; these values are laid 
down in national laws, decrees, or guidance papers (e.g., in Germany: 
BBodSchV 1999). If a threshold value is exceeded, different possibilities 
exist. Risk-reduction measures to decrease the contaminant levels may 
be necessary, and/or the further use of this soil is restricted, because 
the remediation goal for this soil has not been reached. It should be 
evident that those soils which are clearly contaminated, where thresh- 
old values are exceeded, do not have to be tested biologically at all. For 
the other soils, in which such thresholds have not been exceeded, the 
water- extractable ecotoxicological and genotoxic potential is tested. If 
the threshold values of at least one bioassay are exceeded, the source 
of the toxicity should be identified and appropriate measures taken. 
If the test results do not indicate a risk for groundwater or surface 
water, the remediated soil can, for example, be incorporated as sub- 
soil. 

2. If, depending on the envisaged use of the soil, the habitat function 
of the soil has to be assessed, terrestrial ecotoxicological tests have 
to be performed in a second step. Again, if the assessment criteria 
are exceeded, the source of the toxicity should be identified and ap- 
propriate measures taken. If the values are not exceeded, the habitat 
function is substantiated and the remediated soil can be used as top- 
soil. 

In the overview thus far presented it has been shown that ecotoxicological 
test systems are available for the assessment of the retention function and 
for the habitat function of soils. In addition, the results of these tests 
can be evaluated to determine whether the soil might cause a risk to the 
environment. Finally, it should be noted that it may be necessary to modify 



354 



A. Eisentraeger et al. 



Stepl 



Threshold values for chemical 

analysis of contaminants are 

exceeded 



yes 



Limited use of remediated soil - 
remediation target not reached 



no 



Determination of water extractable 

ecotoxicity and genotoxicity: 

threshold values exceeded 



yes 



Toxicity identification 



no 



Low danger of pollutant discharge or 

hazard for groundwater - remediated 

soil can be incorporated as subsoil 



Step 2: (if testing of habitat function is necessary) 



Ecotoxicological testing of habitat 

function of soil: threshold values 

exceeded 



yes 



Toxicity identification 



no 



Habitat function is given 

Remediated soil can be incorporated 
as topsoil 



Fig. 18.5. Stepwise procedure for the examination of soils or soil materials using the test 
systems of Tables 18.1-18.2 and Figs. 18.2-18.4 for remediated soil. (Dechema 2001; Eisen- 
traeger et al. 2004) 



the stepwise procedure presented here in specific cases. These modifications 
might depend on the kind of sample to be tested, the kind of site, the 
kind of contamination, the overall aim of the investigation (precautionary, 
complementary to remediation, on-site analytics), and of course on the 
money available. 



1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 355 

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fahren 



Subject Index 



AAS (atomic absorption spectrometry) 

156, 166 
Abiotic processes 142 
Acclimation 189-197 
Acenaphthene 110 
Acenaphthylen 110 
Acidity 69 

Adenosine diphosphate (ADP) 297 
Adenosine monophosphate (AMP) 297 
Adenosine triphosphate (ATP) 297 
Adenylate energy charge (AEC) 

297-302 
Adenylates 297 

ADP (adenosine diphosphate) 297 
AEC (adenylate energy charge) 

297-302 
Algal growth inhibition test 323, 331 
Aliphatic compounds 100 
Alkanes 103, 189, 274 
Alkyl benzenes 104 
Alkyl naphthalenes 104 
Aluminium (Al) 116 
Ames test 333 
Ammonium 82, 86, 240, 337 
Ammonium oxidation activity 322, 337 
AMP (adenosine monophosphate) 297 
Antagonistic effect 322 
Anthracene 110 
Antibodies 124 
Antimony (Sb) 117 
Apatite 87 

Aromatic hydrocarbons 123, 184 
Arsenic (As) 116 
Atomic absorption spectrometry (AAS) 

156, 166 
ATP (adenosine triphosphate) 297 
Auger 30 

Autochthonous microorganisms 157 
Avena sativa 344 



Barium (Ba) 116 

Benz(a)anthracene 110 

Benzene 101 

Benzene, toluene, ethylbenzene, xylene 

(BTEX) 99, 123, 127, 182, 240, 313 
Benzo(a)pyrene 110 
Benzo(b)fluoranthene 110, 113 
Benzo(ghi)perylene 110 
Benzo(k)fluoranthene 110,113 
Beryllium (Be) 116 
Bioaugmentation 143, 144, 157 
Biodegradation potential 132,189 
Biodegradation rate 98 
Bioleaching 155 
Biological parameters 42, 189, 201, 261, 

281,297,303,309,321 
Bioluminescence 244, 330 
Biomarker 256 
BiomassC 281-288 
Biomass N 289-293 
Biomass, microbial 252,281-295 
Bioreactor experiment 138 
Bioremediation, hydrocarbons 131-153 
Bioremediation, metals 

155-159,161-177 
Bioreporter 233-250 

- systems 235-241 

- MPN analysis 247 

- single point measurement 241 

- vapor phase sensing 244 
Biosurfactant production 145 
Bismuth (Bi) 116 

Blastn 212 

Boron (Bo) 116 

Brassica juncea 168, 173 

Brassica rapa 344 

BTEX (benzene, toluene, ethylbenzene, 

xylene) 99, 123, 127, 182, 240, 313 
Bulk density 52 



362 



Subject Index 



C:N ratio 149 

C:P ratio 149 

Cadmium (Cd) 116,240 

Caesium (Cs) 116 

Calcium (Ca) 116 

Capillary GC-MS 251 

Carbon allocation 190 

Carbon analyzer 288 

Carbon tetrachloride 101 

Catabolic genotypes 208-214 

Catechol-2,3-dioxygenase (xylE) 235 

Cerium (Ce) 116 

Chemical soil properties 71-93 

Chemoautotrophic bacteria 303 

Chlorobenzene 101 

Chloroform 101,240 

Chloroform fumigation 282 

Chromium (Cr) 116,240 

Chrysene 110 

Clod method 57 

Cloning 211-215,226,227 

C0 2 evolution 334 

14 C0 2 evolution 190 

Cobalt (Co) 116,240 

Collembola reproduction test 342 

Colony forming unit (CFU) 273, 278 

Cometabolic degradation 150 

Community structure 209, 218, 254, 255 

Composting 140 

Composting bins 140 

Copper (Cu) 116,240 

Core method 52 

Creosote 109 

Crude oil 179-188,269,274 

CTC (5-cyano-2,3-ditolyl tetrazolium 

chloride) 265 
Culture-dependent methods 201 
Culture-independent methods 201 
Cyano ditolyl tetrazolium chloride (CTC) 

265 
Cycloalkanes 104 
Cyclone fermenters 135 

2,4-D 240 

DAPI (4,6-diamino-2-phenylindole) 265 
Decane 105, 106 
Dehydrogenase activity 316 
Denaturing gradient gel electrophoresis 
(DGGE) 209-224 



Deschampsia caespitosa 173-175 
Desmodesmus subspicatus 331 
DGGE (denaturing gradient gel 

electrophoresis) 209-224 
Diamino phenylindole (DAPI) 265 
Dibenzo(ah)anthracene 110 
Dichlorobenzene 101 
Dichloroethane 101 
Dichloroethene 101 
Dichloromethane 101 
Dichlorophenol 240 
Dichromate oxidation 284 
Diesel 103, 179-188, 310, 313, 316 
Digestion 116 
Dilution 262, 269 
DNA 

- amplification 208 
-polymerase 209,213 

- purification 202, 206 

- quality 207 

- quantity 207 

- sequence identification 216 

- total community 202 
16S rDNA 

- amplification 210 

- cloning and sequencing 211,214 

- genotypes 208 
Drilling 29 

Dry combustion, carbon 72 
Dry combustion, nitrogen 77 
Dry mass 47 

Earthworm mortality 340 

Earthworm reproduction 340 

Ecotoxicity 321-359 

Eicosane 105 

Eisenia andrei 341 

Eisenia fetida 341 

ELISA 121 

EnSys immunoassay 123 

Enumeration, soil microorganisms 261, 

264,268,281-296 
EnviroGard immunoassay 127-130 
Enzyme activity 309-320 
Epifluorescence 264 
Esterase activity 310 
Ethylbenzene 101 
Excavation 32, 54 



Subject Index 



363 



FastA 212,216 
Fatty acids 

- branched unsaturated 254 

- mid-chain branched 254 

- monounsaturated 254 

- naming 257 

- polyunsaturated 254 

- profiles 251-259 

- saturated 254 

- terminally branched 254 
Feasibilibility study 

- bioleaching 155-159 

- hydrocarbons 131-153 

- phy to remediation 161-177 
Fertilization 167 

FID (flame ionization detector) 104 

Field capacity 60 

Field moist 48 

FISH (fluorescent in situ hybridization) 

265 
Flame ionization detector (FID) 104 
Florisil 104 
Fluoranthene 110 
Fluorene 110 

Fluorescein diacetate hydrolysis 313 
Folsomia Candida 343 
Fumigation-extraction 281, 297 
Functional genes 227 
Fungi 155,156,275 

/?-Galactosidase (lacZ) 235 
Gallium (Ga) 116 

Gas chromatography (GC) 100-111, 251 
Gasoline 104, 179-188 
Gasoline-specific compounds 182, 184 
GC (gas chromatography) 100,107,111, 

233 
GC-MS 183 

Gel electrophoresis 204, 206, 207 
Genomics 226 

Genotoxicity 322, 332-334, 349 
Germanium (Ge) 116 
Green fluorescent protein (GFP) 236 
Growth-inhibition assay 331 

Habitat function 322, 334, 350 
Halogenated hydrocarbons 101 
Headspace analyzer 100 
Heating oil 103 



Heavy metal: see metal 
Heterotrophic leaching 155 
High-performance liquid chromatography 

(HPLC) 297 
Hopane 179-188 
HPLC (high-performance liquid 

chromatography) 297 
Humic acids 71, 203 
Humification 72 
Humus 71 

Hydrindantin (ninhydrin) 289 
Hydrocarbons 

- biodegradation 179-199 

- 14 C-labeled 189 

- degrading bacteria 142 

- mineralization 196 

- monoaromatic 100, 189 

- polycyclic aromatic 106, 109, 123, 127, 
189,310,313,316 

- substrates 191,268,275-277 

- volatile 99, 184, 195, 268, 277 
Hydrolases, hydrolytic activity 310,313 

ICP-AES (inductively-coupled 

plasma-atomic emission spectrometry) 

156, 166 
Immobilization 161, 171 
Immunoassays 121-130 

- detection limits 123 

- ELISA 121 

- EnSys 123 

- EnviroGard 127-130 

- RaPID 121-126 
Indeno(l,2,3-cd)pyrene 110 
Indium (In) 116 
Inoculum 134, 157 

INT (iodonitrotetrazolium chloride) 317 

Internal markers 179 

Internal standards 101, 105, 112, 116, 182 

Iodonitrotetrazolium chloride (INT) 317 

Iron(Fe) 116,240 

Isoalkanes 104 

Isoparaffins 184 

Jet fuel 103,274 

Kerosene 99 
Kjeldahl method 79 



364 



Subject Index 



fi-Lactamase {bid) 236 

Land treatment 142 

Lead(Pb) 116,240 

Lipase activity 310 

Liquid scintillation counter 191 

Lithium (Li) 116 

Loss on ignition (LOI) 72, 74 

Lubrication oil 103 

Luciferase AB (luxAB) 239 

Luciferase CDABE (luxCDABE) 239 

Luciferase, bacterial (lux) 238 

Luciferase, insect (luc) 238 

Luminescence inhibition assay 330 

Magnesium (Mg) 116 

Mangan (Mn) 116 

Mass selective detector (MSD) 102, 1 1 1 

Matric pressure 60 

Maximum water-holding capacity 5 1 

Membrane filter 265 

Mercury (Hg) 116,240 

Metagenomic libraries 226 

Metal 115-118, 155-159, 161-177 

- accumulation 168 

- immobilization 155, 161 

- leaching 155 

- mobilization 155 

- speciation 115 
Metal-contaminated soil 155, 161 
Methyl tert-butyl ether (MTBE) 99 
Microarrays 227 

Microbial activity 1 94, 309 
Microbial community 209, 218, 254, 255 
Microbial enrichment, selective 134 
Microbial remediation 131-153, 155-159 
Microscope, enumeration 264 
Microtiter plate 270 
Microtox 239 
Microwave oven 116 
Mineralization 72, 189 
Mineralization potential 193 
Mini-transposon 239 
Molecular techniques 202-231 
Monitoring 98, 233, 309 

- biodegradation of hydrocarbons 142, 
310,313,316 

- biodegradation of carboxyl esters 310 



- biodegradation of surfactants 316 

- impact of contamination 321 

- impact of fertilization 197,313 

- of process 142 

Monoaromatic hydrocarbons 100, 189 
Most probable number (MPN) 268, 247 
MPN (most probable number) 268, 247 
MTBE (methyl tert-butyl ether) 99 

Naphthalene 100, 110, 182, 240, 274 
Nickel (Ni) 116,240 
Ninhydrin (hydrintantin) 289 
Ninhydrin-reactive nitrogen 

289-291 
Niobium (Nb) 116 
Nitrate 82, 84, 240, 337 
Nitrification 303, 337 
Nitrite 82, 337 
Nitrobacter 303 
Nitrogen (N) 

- immobilization 306 

- inorganic 82 

- mineralization 303-308 

- ninhydrin-reactive 289-291 

- organic 303 

- total 76,292 
Nitrosomonas 303 
Nutrient sources 149, 156 
Nutritional factors 148 

O2 consumption 335 
Oligonucleotide sequences 212 
Organic acids 156, 158 
Organic carbon 71 
Organic matter 71 
Organic nitrogen 303 
Oven oxidation 288 
Overlayer plate 274, 276 
Oxido reductases 317 

PAHs (polycyclic aromatic hydrocarbons) 
106, 109, 123, 127, 189, 310, 313, 316 

- alkylated 184 

- deuterated 110 

- native 110 
Paraffins 184 

PCBs (polychlorinated biphenyls) 127, 
208, 240 



Subject Index 



365 



PCR amplification 208-22 1 

- catabolic genotypes 210,213 

- control 214 

- 16SrDNA 210 

PCR cloning kits 210,215 
PCR primers 

- catabolic genes 212 

- 16SrDNA 214 

- universal 220 

PCR thermocycler 210 

Pentatriacontane 105 

Percolation 156, 157 

Percussion boring 28 

Permeability 48, 52 

Petroleum hydrocarbons 98, 123, 127, 

137,139,142,184,189,310,316 
pH value 68 
Phenanthrene 1 10, 274 
Phenol 240 
Phosphorus (P) 

- labile 90 

- quantification 91, 116 

- total 88 

Physical soil properties 47-71 

Phytoextraction 161, 163, 168, 172 

Phytoremediation 161 

Phytostabilization 161, 171-175 

Plant growth test 344 

Plants 161 

Plate count 272 

Platinum catalyzer 288 

PLFA (polar lipid fatty acids) 251, 253 

Polar lipid fatty acids (PLFA) 251, 253 

Polychlorinated biphenyls (PCBs) 127, 

208, 240 
Polycyclic aromatic hydrocarbons (PAHs) 

106, 109, 123, 127, 189, 310, 313, 316 
Polyvinylpolypyrrolidone (PVPP) 206 
Pore size distribution 59 
Pore water pressure 50 
Porosity 52 
Potassium (K) 116 
Pour plate 276 
Pre-extraction 284, 293 
Pre-incubation 298 
Pressure plate extractor 65 
Primer design 212 



Priority pollutants 181 

Proton activity 68 

Purge and trap 182 

PVPP (polyvinylpolypyrrolidone) 206 

Pyrene 110 

Radioactivity 198 

Radiolabeled substrates 269 

Radiorespirometry 189 

RaPID Immunoassay 121-126 

Redox titration 284 

Reporter systems 235 

Respiration activity 193 

Respiration curve test 334 

Respirometer 136 

Retention time 102 

Rhenium (Re) 116 

Ribosomal Database Project (RDP) 217 

Rubidium (Rb) 116 

Salmonella typhimurium 333 
Salmonella/microsome test (Ames test) 

323, 333 
Scale up 141 
Scintillation cocktail 192 
Segmented flow analysis (SFA) 85 
Selected ion monitoring (SIM) 184 
Selective microbial enrichment 134 
Selenium (Se) 116 
Sequencing 211-215 
Serial dilution 262, 263, 270 
Sewage sludge 140 
SFA (segmented flow analysis) 85 
Shotgun cloning 226, 227 
Silicon (Si) 116 
Silver (Ag) 116,240 
SIM (selected ion monitoring) 184 
Slurry bioreactors 137 
Sodium (Na) 116 
Soil 

- aeration 52 

- aggregates 264 

- biological activity 309 

- column 134 

- DNA: see DNA 

- enzymes 309 

- microcosm 136 

- nucleic acid extraction 202, 205 



366 



Subject Index 



- nutrients 76 

- pores 59 

- quality 2 

Soil sampling 1-37, 132-134 

- along a linear source 21 

- circular grids 15 

- container 34 

- equipment 133 

- irregular sampling 12 

- methods 25-37 

- non- systematic patterns 12 

- pretreatment 37-40 

- quantity 24 

- random sampling 17 

- rectangular grid 20 

- regular grids 16 

- sample type 25 

- sampler 32 

- sampling points 10 

- sampling site 22 

- strategy 7-25 

- stratified random sampling 17 

- systematic sampling 16 

- unaligned random sampling 19 

- undisturbed samples 27 
Soil storage 41-44 

Soil water characteristics 62, 65 
Solid phase extraction 329 
Solid-phase microextraction (SPME) 

143 
Solvent extraction 183 
SOS Chromotest 235 
SPME (solid-phase microextraction) 
Spore suspension 158 
Spray plate 276 
Spread plate 275 
Sterile control 135 
Streamline test 166 
Stripping apparatus 192 
Strontium (Sr) 116 
Styrene 101, 182 
Suction table 62 
Sulfur (S) 116 
Surfactants 144-147 
Synergistic effect 322 



143 



Tellurium (Te) 116 

Tert-amyl methyl ether (TAME) 99 

Tetrachloroethene 101 

Tetracontane 105, 106 

Tetramethylbutane 179 

Threshold values 346 

Time domain reflectometry (TDR) 48 

Titanium (Ti) 116 

Toluene 101,274 

Toxicity 321-359 

TPHs (total petroleum hydrocarbons) 98, 

127, 137, 139, 142, 184, 189, 310, 316 
Transposon 239 
Treatability study 162-166 

- bioleaching 155-159 

- hydrocarbons 131-153 

- phy to remediation 161-177 
Triacontane 105 
Trichlorobenzene 101 
Trichloroethane 101 
Trichlorophenol 240 
Trimethyl cyclopentane 179 
Trimethyl phenanthrene 179 
Tungsten (W) 116 



Umu test 322, 332 

Uranium (U) 116 

Urogen III methyltransferase (UMT) 

UV-persulfate oxidation 286 



237 



Vanadium (V) 116 
Vapor phase contaminants 244 
Vapor plate 277 
Vibrio fischeri 323, 330 
Vitotox 239 

Volatile hydrocarbons 99, 184, 195, 268, 
277 



Water content 47 
Water content adjustment 
Water extraction 329 
Water retention 59 
Water-holding capacity 50 
Wilting point 60 

Xylene 101,182,240,274 



51 



TAME (tert-amyl methyl ether) 99 
Tantalum (Ta) 116 



Zinc(Zn) 116,240 
Zirconium (Zr) 116